Method to determine TGF-β

ABSTRACT

A method for treating or preventing cardiovascular pathologies by administering a compound of the formula (I):                    
     wherein Z is C═O or a covalent bond; Y is H or O(C 1 -C 4 )alkyl, R 1  and R 2  are individually (C 1 -C 4 )alkyl or together with N are a saturated heterocyclic group, R 3  is ethyl or chloroethyl, R 4  is H or together with R 3  is —CH 2 —CH 2 — or —S—, R 5  is I, O(C 1 -C 4 )alkyl or H and R 6  is I, O(C 1 -C 4 )alkyl or H with the proviso that when R 4 , R 5 , and R 6  are H, R 3  is not ethyl; or a pharmaceutically acceptable salt thereof, effective to activate or stimulate production of TGF-beta to treat and/or prevent conditions such as atherosclerosis, thrombosis, myocardial infarction, and stroke is provided. Useful compounds include idoxifene and salts thereof. Further provided is a method for identifying a compound that is a TGF-beta activator or production stimulator is provided. Another embodiment of the invention is an assay or kit to determine TGF-beta in vitro. Also provided is a therapeutic method comprising inhibiting smooth muscle cell proliferation associated with procedural vascular trauma employing the administration of tamoxifen or structural analogs thereof, including compounds of formula (I).

RELATED APPLICATIONS

This application is a continuation-in-part of U.S. Ser. No. 08/242,161,filed May 12, 1994, now U.S. Pat. No. 5,847,007, which is acontinuation-in-part of U.S. Ser. No. 08/061,714, filed May 13, 1993,abandoned, which are incorporated by reference herein. This applicationis also a continuation-in-part of U.S. Ser. No. 08/241,844, filed May12, 1994, abandoned, which is a continuation-in-part of U.S. Ser. No.08/062,451, filed May 13, 1993, abandoned which are incorporated byreference herein.

FIELD OF THE INVENTION

This invention relates generally to the prevention and treatment ofcardiovascular pathologies. More specifically, a method for treating orpreventing atherosclerosis is provided.

BACKGROUND OF THE INVENTION

Many pathological conditions have been found to be associated withsmooth muscle cell proliferation. Such conditions include restenosis,atherosclerosis, coronary heart disease, thrombosis, myocardialinfarction, stroke, smooth muscle neoplasms such as leiomyoma andleiomyosarcoma of the bowel and uterus, uterine fibroid or fibroma, andobliterative disease of vascular grafts and transplanted organs. Themechanisms of abnormal smooth muscle cell proliferation are not yet wellunderstood.

For example, percutaneous transluminal coronary angioplasty (PTCA) iswidely used as the primary treatment modality in many patients withcoronary artery disease. PTCA can relieve myocardial ischemia inpatients with coronary artery disease by reducing lumen obstruction andimproving coronary flow. The use of this surgical procedure has grownrapidly, with 39,000 procedures performed in 1983, nearly 150,000 in1987, 200,000 in 1988, 250,000 in 1989, and over 500,000 PTCAs per yearare estimated by 1994. Stenosis following PTCA remains a significantproblem, with from 25% to 35% of the patients developing restenosiswithin 1 to 3 months. Restenosis results in significant morbidity andmortality and frequently necessitates further interventions such asrepeat angioplasty or coronary bypass surgery. No surgical interventionor post-surgical treatment (to date) has proven effective in preventingrestenosis.

The processes responsible for stenosis after PTCA are not completelyunderstood but may result from a complex interplay among severaldifferent biologic agents and pathways. Viewed in histological sections,restenotic lesions may have an overgrowth of smooth muscle cells in theintimal layers of the vessel. Several possible mechanisms for smoothmuscle cell proliferation after PTCA have been suggested. For example,Barath et al. (U.S. Pat. No. 5,242,397) disclose delivering cytotoxicdoses of protein kinase C inhibitors, including tamoxifen, locally bycatheter to the site of the atherosclerotic lesion.

Compounds that reportedly suppress smooth muscle proliferation in vitromay have undesirable pharmacological side effects when used in vivo.Heparin is an example of one such compound, which reportedly inhibitssmooth muscle cell proliferation in vitro but when used in vivo has thepotential adverse side effect of inhibiting coagulation. Low molecularweight fragments of heparin, while having reduced anti-coagulantactivity, have the undesirable pharmacological property of a shortpharmacological half-life. Attempts have been made to solve suchproblems by using a double balloon catheter, i.e., for regional deliveryof the therapeutic agent at the angioplasty site (e.g., U.S. Pat. No.4,824,436), and by using biodegradable materials impregnated with adrug, i.e., to compensate for problems of short half-life (e.g., U.S.Pat. No. 4,929,602).

In general, atherosclerosis is a cardiovascular disease in which thevessel wall is remodeled, compromising the lumen of the vessel. Theatherosclerotic remodeling process involves accumulation of cells, bothsmooth muscle cells and monocyte/macrophage inflammatory cells, in theintima of the vessel wall. These cells take up lipid, likely from thecirculation, to form a mature atherosclerotic lesion. Although theformation of these lesions is a chronic process, occurring over decadesof an adult human life, the majority of the morbidity associated withatherosclerosis occurs when a lesion ruptures, releasing thrombogenicdebris that rapidly occludes the artery. When such an acute event occursin the coronary artery, myocardial infarction can ensue, and in theworst case, can result in death.

The formation of the atherosclerotic lesion can be considered to occurin five overlapping stages such as migration, lipid accumulation,recruitment of inflammatory cells, proliferation of vascular smoothmuscle cells, and extracellular matrix deposition. Each of theseprocesses can be shown to occur in man and in animal models ofatherosclerosis, but the relative contribution of each to the pathologyand clinical significance of the lesion is unclear.

Thus, a need exists for therapeutic methods and agents to treatcardiovascular pathologies, such as atherosclerosis and other conditionsrelated to coronary artery disease.

SUMMARY OF THE INVENTION

A therapeutic method for preventing or treating a cardiovascularindication characterized by a decreased lumen diameter is provided. Themethod comprises administering to a mammal at risk of, or afflictedwith, said cardiovascular indication, a cytostatic dose of a TGF-betaactivator or production stimulator. The cytostatic dose is effective toactivate or stimulate production of TGF-beta and the effective amountinhibits smooth muscle cell proliferation, inhibits lipid accumulation,increases plaque stability, or any combination thereof.

A therapeutic method is provided for treating or preventingcardiovascular pathologies, such as conditions selected from the groupconsisting of atherosclerosis, thrombosis, myocardial infarction, andstroke. The method comprises the systemic or local administration of anamount of a compound of formula (I)

wherein Z is C═O or a covalent bond; Y is H or O(C₁-C₄)alkyl, R¹ and R²are individually (C₁-C₄)alkyl or together with N are a saturatedheterocyclic group, R³ is ethyl or chloroethyl, R⁴ is H or together withR³ is —CH₂—CH₂— or —S—, R⁵ is I, O(C₁-C₄)alkyl or H and R⁶ is I,O(C₁-C₄)alkyl or H with the proviso that when R⁴, R⁵, and R⁶ are H, R³is not ethyl; or a pharmaceutically acceptable salt, including mixturesthereof, effective to activate or stimulate production of TGF-beta in amammal afflicted with one of these conditions. Thus, in this embodimentof the invention, the compound of formula (I) does not includetamoxifen.

The administered compound of formula (I) can act on vascular smoothmuscle cells (VSMC) to inhibit the pathological activity of these smoothmuscle cells and can inhibit lipid proliferative lesions. Preferably,the compound significantly reduces the rate of completion of the cellcycle and cell division, and preferably is administered at cytostatic,as opposed to cytotoxic, doses. A preferred embodiment of the inventioncomprises treatment of atherosclerosis, wherein the compound of formula(I), such as idoxifene or idoxifene salt, inhibits lipid accumulation byvascular smooth muscle cells and/or stabilizes an arterial lesionassociated with atherosclerosis, i.e., increases plaque stability, toprevent rupture or growth of the lesion. As exemplified hereinbelow,orally administered tamoxifen significantly inhibits the formation oflipid lesions, induced by a high fat diet, in C57B16 mice and in thetransgenic apo(a) mouse. The 90% reduction in lesion area and number inboth of these mouse models indicates that tamoxifen affects theaccumulation of lipid in the cells and stroma of the vessel wall. Theinhibition of lipid accumulation and lesion development in these treatedmice indicates that tamoxifen and analogs thereof, as well as compoundsof formula (I), may inhibit the development of atherosclerotic lesionsin humans by inhibiting lipid accumulation, in addition to decreasingsmooth muscle cell proliferation.

Other preferred embodiments of the invention comprise the localadministration of the compound of formula (I) to an arterial lesionassociated with atherosclerosis, and a kit to accomplish saidadministration.

A further embodiment of the invention is a method for preventingcardiovascular pathologies in a mammal at risk of such a condition. Suchconditions include atherosclerosis, thrombosis, myocardial infarction,and stroke. The method comprises the administration of an amount of thecompound of formula (I) to a mammal, such as a human, effective toactivate or stimulate production of TGF-beta. The amount of the compoundis administered over time as a preventative measure. Preferably, thecompound is administered orally, in a series of spaced doses.

A further embodiment of the invention is a method for inhibiting smoothmuscle cell (SMC) proliferation associated with procedural vasculartrauma as by the systemic or localized catheter or non-catheteradministration to a mammal, such as a human patient, subjected to saidprocedure, an effective cytostatic SMC proliferation inhibitory amountof tamoxifen (TMX), a compound of formula (I), a combination thereof, ora pharmaceutically acceptable salt thereof. The systemic administrationcan be accomplished by oral or parenteral administration of one of moresuitable unit dosage forms, which, as discussed below, may be formulatedfor sustained release. The administration may be essentially continuousover a preselected period of time or may be in a series of spaced doses,either before, during, or after the procedural vascular trauma, or bothbefore and after the procedural trauma, including during the procedurecausing the trauma.

As used herein, the term “procedural vascular trauma” includes theeffects of surgical/mechanical interventions into mammalian vasculature,but does not include vascular trauma due to the organic vascularpathologies listed hereinabove.

Thus, procedural vascular traumas within the scope of the presenttreatment method include (1) organ transplantation, such as heart,kidney, liver and the like, e.g., involving vessel anastomosis; (2)vascular surgery, such as coronary bypass surgery, biopsy, heart valvereplacement, atheroectomy, thrombectomy, and the like; (3) transcathetervascular therapies (TVT) including angioplasty, e.g., laser angioplastyand PTCA procedures discussed hereinbelow, employing balloon catheters,and indwelling catheters; (4) vascular grafting using natural orsynthetic materials, such as in saphenous vein coronary bypass grafts,dacron and venous grafts used for peripheral arterial reconstruction,etc.; (5) placement of a mechanical shunt, such as a PTFE hemodialysisshunt used for arteriovenous communications; and (6) placement of anintravascular stent, which may be metallic, plastic or a biodegradablepolymer. See U.S. patent application Ser. No. 08/389,712, filed Feb. 15,1995, which is incorporated by reference herein. For a generaldiscussion of implantable devices and biomaterials from which they canbe formed, see H. Kambic et al., “Biomaterials in Artificial Organs”,Chem. Eng. News, (Apr. 14, 1986), the disclosure of which isincorporated by reference herein.

In the case of organ transplantation, the entire organ, or a portionthereof, may be infused with a solution of TMX and/or the compound offormula (I), prior to implantation. Likewise, in vascular surgery, thetermini of the vessels subject to anastomosis can be infused with TMXand/or the compound of formula (I), or the antiproliferative agents canbe delivered from pretreated sutures or staples.

The delivery of TGF-beta activators or production stimulators to thelumen of a vessel via catheter, before, during or after angioplasty, isdiscussed in detail below. A stent or shunt useful in the present methodcan comprise a biodegradable coating or porous non-biodegradablecoating, having dispersed therein the sustained-release dosage form. Inthe alternative embodiment, a biodegradable stent or shunt may also havethe therapeutic agent impregnated therein, i.e., in the stent or shuntmatrix. Utilization of a biodegradable stent or shunt with thetherapeutic agent impregnated therein is further coated with abiodegradable coating or with a porous non-biodegradable coating havingthe sustained release-dosage form dispersed therein is alsocontemplated. This embodiment of the invention would provide adifferential release rate of the therapeutic agent, i.e., there would bea faster release of the therapeutic agent from the coating followed bydelayed release of the therapeutic agent that was impregnated in thestent or shunt matrix upon degradation of the stent or shunt matrix. Theintravascular stent or shunt thus provides a mechanical means ofmaintaining or providing an increase in luminal area of a vessel, andthe antiproliferative agent inhibits the VSMC proliferative responseinduced by the stent or shunt, which can cause occlusion of blood flowand coronary failure.

For local administration during grafting, the ex vivo infusion of theantiproliferative agent into the excised vessels (arteries or veins) tobe used for vascular grafts can be accomplished. In this aspect of theinvention, the vessel that is to serve as the graft is excised orisolated and subsequently distended by an infusion of a solution of thetherapeutic agent, preferably by pressure infusion. Of course, grafts ofsynthetic fiber can be precoated with TMX and/or compounds of formula(I) prior to in vivo placement.

A further aspect of the invention is a method comprising inhibitingvascular smooth muscle cell proliferation associated with proceduralvascular trauma due to organ transplantation, vascular surgery,angioplasty, shunt placement, stent placement or vascular graftingcomprising administration to a mammal, such as a human, subjected tosaid procedural trauma an effective antiproliferative amount of acompound of formula (I) or a pharmaceutically acceptable salt thereof.Administration may be systemic, as by oral or parenteral administration,or local, as to the site of the vascular trauma, or both.

Yet a further aspect of the invention provides a method comprisinginhibiting non-aortal vascular smooth muscle cell proliferationassociated with procedural vascular trauma comprising administering aneffective cytostatic antiproliferative amount of tamoxifen or astructural analog thereof, including the pharmaceutically acceptablesalts thereof, to a mammal, such as a human, subjected to saidprocedural vascular trauma. Said administration can be systemic or bylocal, catheter or non-catheter delivery to the site of the trauma.

Also provided is a kit comprising packing material enclosing, separatelypackaged, a catheter, a stent, a shunt or a synthetic graft and a unitdosage form of an amount of a compound of formula (I) and/or tamoxifeneffective to accomplish these therapeutic results when deliveredlocally, as well as instruction means for its use, in accord with thepresent methods.

Another embodiment of the present invention is a method for identifyinga compound which is a TGF-beta activator or production stimulator. Humanvascular smooth muscle cells (hVSMC) are cultured with an amount of thecompound effective to reduce the normal rate of hVSMC proliferation, dueto TGF-beta activation or production stimulation by said compound. Thenthe hVSMC are contacted with an amount of an antibody which neutralizesTGF-beta activity. The method can also include the culture of rat aorticvascular smooth muscle cells (rVSMC) with an amount of the same compoundeffective to reduce the normal rate of proliferation of rVSMC, due toTGF-beta activation or production stimulation by said compound. TherVSMC are then contacted with the neutralizing antibody. The restorationof a normal rate of proliferation in treated rVSMC and treated hVSMCafter contact with the TGF-beta neutralizing antibody indicates that thereduction of proliferation is due to TGF-beta activation or productionstimulation in rVSMC and hVSMC by said compound, and suggests that hVSMCwould be amenable to treatment by the administration of said compound invivo.

Useful compounds of formula (I) are TGF-beta activators and TGF-betaproduction stimulators. These compounds, including their salts andmixtures thereof, may be employed in the practice of the presentinvention to prevent or treat other conditions characterized byinappropriate or pathological activity of vascular smooth muscle cells.Such TGF-beta activators and production stimulators inhibit abnormalactivity of vascular smooth muscle cells. Preferred compounds of formula(I) include those wherein Z is a covalent bond, Y is H, R³ is ClCH₂CH₂or ethyl, R⁵ or R⁶ is iodo, R⁴ is H or with R³ is —CH₂CH₂— or —S—, R¹and R² are each CH₃ or together with N are pyrrolidino,hexamethyleneimino or piperidino. These compounds can include structuralanalogs of tamoxifen (including derivatives of TMX and derivatives ofsaid analogs) having equivalent bioactivity. Such analogs includeidoxifene(IDX)(E-1-[4-[2-N-pyrrolidino)ethoxy]phenyl]-1-(4-iodophenyl)-2-phenyl-1-butene),raloxifene, 3-iodotamoxifen, 4-iodotamoxifen, tomremifene, and thepharmaceutically acceptable salts thereof.

Also provided are a method and a kit to determine the presence andamount of TGF-beta in a sample containing TGF-beta. The method for thedetermination of TGF-beta in vitro can be used to identify a patient atrisk for atherosclerosis and/or monitor a recipient that has receivedone or more administrations of a TGF-beta activator or productionstimulator. Blood serum or plasma or tissue from a patient or recipientis contacted with a capture moiety to form a capture complex of saidcapture moiety and TGF-beta. Preferably, the capture moiety is animmobilized capture moiety. The capture complex is then contacted with adetection moiety capable of binding TGF-beta comprising a detectablelabel, or a binding site for a detectable label, to form a detectablecomplex. The presence and amount, or absence, of the detectable complexis then determined, thereby determining the presence and amount, orabsence, of TGF-beta in the blood of the patient or recipient.

A test kit for determining TGF-beta in vitro includes packaging materialenclosing (a) a capture moiety capable of binding TGF-beta, and (b) adetection moiety capable of binding to TGF-beta, where the detectionmoiety has a detectable label or a binding site for a detectable label.The capture moiety and the detection moiety are separately packaged inthe test kit. Preferably, the capture moiety is solidsubstrate-immobilized. Preferably, the capture moiety is the TGF-betatype II receptor extracellular domain. More preferably, the TGF-betatype II receptor extracellular domain is derived from a bacterialexpression system. The kit can also comprise instruction means forcorrelation of the detection or determination of TGF-beta with theidentification of the patients or monitoring discussed above.

Further provided is a method for upregulating cellular mRNA coding forTGF-beta. Cells (e.g., smooth muscle cells) amenable to suchmanipulation of mRNA accumulation are identified in the manner describedherein and are exposed to an effective amount of a TGF-beta mRNAregulator (i.e., a subset of TGF-beta production stimulators), eitherfree or in a sustained-release dosage form. In this manner, TGF-betaproduction is stimulated.

In addition, methods for using TGF-beta to maintain and increase vessellumen diameter in a diseased or injured mammalian vessel are described.

DESCRIPTION OF THE DRAWINGS

FIGS. 1 and 2 depict pathways for the modulation of vascular smoothmuscle cell proliferation in vivo.

DETAILED DESCRIPTION OF THE INVENTION

As used herein the following terms have the meanings as set forth below:

“Proliferation,” means an increase in cell number, i.e., by mitosis ofthe cells.

“Abnormal or Pathological or Inappropriate Activity or Proliferation”means division, growth or migration of cells occurring more rapidly orto a significantly greater extent than typically occurs in a normallyfunctioning cell of the same type, or in lesions not found in healthytissues.

“Expressed” means mRNA transcription and translation with resultantsynthesis, glycosylation, and/or secretion of a polypeptide by a cell,e.g., chondroitin sulfate proteoglycan (CSPG) synthesized by a vascularsmooth muscle cell or pericyte.

The term “tamoxifen”, as used herein, includestrans-2-[4-(1,2-diphenyl-1-butenyl)phenoxy]-N,N-dimethylethylamine, andthe pharmaceutically acceptable salts thereof, which are capable ofenhancing the production or activation of TGF-beta. The activated formof TGF-beta, in turn, inhibits vascular smooth muscle cell activity.Isomers and derivatives of the aforementioned chemical compound are alsoincluded within the scope of the term “tamoxifen” for the purposes ofthis disclosure.

The term “structural analogs thereof” with respect to tamoxifenincludes, but is not limited to, all of the compounds of formula (I)which are capable of enhancing production or activation of TGF-beta.See, for example, U.S. Pat. No. 4,536,516, and U.K. Patent 1,064,629.

Because tamoxifen (TMX) causes liver carcinogenicity in rats and hasbeen correlated with an increased risk of endometrial cancer in womenand may increase the risk of certain gut cancers, other tamoxifenanalogs may be considered safer to administer if they are lesscarcinogenic. The carcinogenicity of TMX has been attributed to theformation of covalent DNA adducts. Of the TMX analogs and derivatives,only TMX and toremifene have been studied for long-term carcinogenicityin rats and these studies provide strong evidence that covalent DNAadducts are involved in rodent hepatocarcinogenicity of TMX. Toremifene,which exhibits only a very low level of hepatic DNA adducts, was foundto be non-carcinogenic. See Potter et al., Carcinogenesis, 15, 439(1994).

It is postulated that 4-hydroxylation of TMX yields electrophilicalkylating agents which alkylate DNA through the ethyl group of TMX.This mechanistic hypothesis explains the low level of DNA adductformation by the non-TMX analogs of formula (I), including the TMXanalog toremifene and the absence of DNA adducts detected for theanalogs 4-iodotamoxifen and idoxifene. Thus, all of these analogs arelikely to be free from the risk of carcinogenesis in long term use. SeePotter et al., supra. Idoxifene includes(E)-1-[4-[2-(N-pyrrolidino)ethoxy]phenyl]-1-(4-iodophenyl)-2-phenyl-1-buteneand its pharmaceutically acceptable salts and derivatives. See R.McCague et al., Organic Preparations and Procedures Int., 26, 343 (1994)and S. K. Chandler et al., Cancer Res., 51, 5851 (1991). Besides itslower potential for inducing carcinogenesis via formation of DNA adductswhich can damage DNA, other advantages of IDX compared with TMX are thatIDX has reduced residual oestrogenic activity in rats and an improvedmetabolic profile. IDX is the preferred embodiment of the presentinvention.

Also included within the scope of the term tamoxifen are the TMXstructural analogs toremifene and raloxifene, metabolites orpharmaceutically acceptable salts thereof. Other “antisteroids” or“steroidal antagonists” can also be useful as TGF-beta activators orproduction stimulators or lead compounds, including other knownstilbene-type. antisteroids including cis- and trans-clomiphene,droloxifene,(1-[4-(2-dimethylaminoethoxy)phenyl]-1-(3-hydroxyphenyl)-2-phenyl-2-butene(see U.S. Pat. No. 5,384,332), 1-nitro-1-phenyl-2-(4-hydroxyphenyl oranisyl)-2-[4-(2-pyrrol-N-ylethoxy)-phenyl]ethylene(CN-55,945),trans-1,2-dimethyl-1,2-(4-hydroxyphenyl)ethylene(trans-dimethylstilboestrol),trans-diethylstilboestrol, and1-nitro-1-phenyl-2-(4-hydroxyphenyl)-2-[4-(3-dimethylaminopropyloxy)phenyl-ethylene(GI680).

Known 1,2-diphenylethane-type antisteroids includecis-1,2-anisyl-1-[4-(2-diethylaminoethoxy)phenyl]ethane (MRL-37),1-(4-chlorophenyl) 1-[4-(2-diethylaminoethoxy)phenyl]-2-phenylethanol(WSM-4613); 1-phenyl-1-[4-(2-diethylaminoethoxy)phenyl]-2-anisylethanol(MER-25); 1-phenyl-1-[4-(2-diethylaminoethoxy)phenyl)-2-anisyl-ethane,mesobutoestrol (trans-1,2-dimethyl-1,2-(4-hydroxyphenyl)-ethane),meso-hexestrol, (+)hexestrol and (−)-hexestrol.

Known naphthalene-type antisteroids include nafoxidine,1-[4-(2,3-dihydroxypropoxy)phenyl]-2-phenyl-6-hydroxy-1,2,3,4-tetrahydro-naphthalene,1-(4-hydroxyphenyl)-2-phenyl-6-hydroxy-1,2,3,4-tetrahydronaphthalene,1-[4-(2-pyrrol-N-ylethoxy)-phenyl]-2-phenyl-6-methoxy-3,4-dihydronaphthalene(U11, 100A), and1-[4-(2,3-dihydroxypropoxy)phenyl]-2-phenyl-6-methoxy-3,4-dihydronaphthalene(U-23, 469).

Known antisteroids which do not fall anywhere within these structuralclassifications include coumetstrol, biochanin-A, genistein,methallenstril, phenocyctin, and1-[4-(2-dimethylaminoethoxy)phenyl]-2-phenyl-5-methoxyindene (U, 11555).In the nomenclature employed hereinabove, the term “anisyl” is intendedto refer to a 4-methoxyphenyl group.

The pharmaceutically acceptable inorganic and organic acid amine saltsof the amino group-containing antisteroids are also included within thescope of the term “antisteroid”, as used herein, and include citrates,tartrates, acetates, hydrochlorides, hydrosulfates and the like.

“TGF-beta” includes transforming growth factor-beta as well asfunctional equivalents, derivatives and analogs thereof. The TGF-betaisoforms are a family of multifunctional, disulfide-linked dimericpolypeptides that affect activity, proliferation and differentiation ofvarious cells types. TGF-beta is a polypeptide produced in a latentpropeptide form having, at this time, no identified biological activity.To be rendered active and, therefore, capable of inhibiting vascularsmooth muscle cell proliferation, the propeptide form of TGF-beta mustbe cleaved to yield active TGF-beta.

“TGF-beta activator” includes moieties capable of directly or indirectlyactivating the latent form of TGF-beta to the active form thereof. Anumber of the compounds of formula (I) are believed to be TGF-betaactivators.

“TGF-beta production stimulator” includes moieties capable of directlyor indirectly stimulating the production of TGF-beta (generally thelatent form thereof). Such TGF-beta production stimulators may beTGF-beta mRNA regulators (i.e., moieties that increase the production ofTGF-beta mRNA), enhancers of TGF-beta mRNA expression or the like.

“Direct” action implies that the TGF-beta activator acts on the latentform of TGF-beta. Such direct action, when applied to TGF-betaproduction stimulators, indicates that cells upon which the productionstimulator acts increase TGF-beta mRNA production or expression ofTGF-beta.

“Indirect” action implies that the TGF-beta activator acts on a moietythat itself or through one or more other moieties acts on latentTGF-beta. Such indirect action, when applied to TGF-beta productionstimulators, indicates that the stimulators act on a moiety that itselfor through one or more other moieties acts on a population of cells tostimulate the production of TGF-beta mRNA or the expression of TGF-beta.

“Sustained release” means a dosage form designed to release atherapeutic agent therefrom for a time period ranging from about 3 toabout 21 days. Release over a longer time period is also contemplated asa “sustained release” dosage form of the present invention.

For the purposes of this description, the prototypical cells, upon whichthe effects of TGF-beta activators or production stimulators are felt,are smooth muscle cells and pericytes derived from the medial layers ofvessels which proliferate in intimal hyperplastic vascular sitesfollowing injury, such as that caused during PTCA. TGF-beta activatorsand production stimulators are not restricted in use for therapyfollowing angioplasty; rather, the usefulness thereof will be proscribedby their ability to inhibit abnormal cellular proliferation, forexample, of smooth muscle cells and pericytes in the vascular wall.Thus, other aspects of the invention include TGF-beta activators orproduction stimulators used in early therapeutic intervention forreducing, delaying, or eliminating (and even reversing) atheroscleroticplaque formation and areas of vascular wall hypertrophy and/orhyperplasia. TGF-beta activators and production stimulators also findutility for early intervention in pre-atherosclerotic conditions, e.g.,they are useful in patients at a high risk of developing atherosclerosisor with signs of hypertension resulting from atherosclerotic changes invessels or vessel stenosis due to hypertrophy of the vessel wall.

TGF-beta activators or production stimulators of the invention areuseful for inhibiting the pathological proliferation of vascular smoothmuscle cells, e.g., for reducing, delaying, or eliminating stenosisfollowing angioplasty. As used herein the term “reducing” meansdecreasing the intimal thickening that results from stimulation ofsmooth muscle cell proliferation following angioplasty, either in ananimal model or in man. “Delaying” means delaying the time until onsetof visible intimal hyperplasia (e.g., observed histologically or byangiographic examination) following angioplasty and may also beaccompanied by “reduced” restenosis. “Eliminating” restenosis followingangioplasty means completely “reducing” intimal thickening and/orcompletely “delaying” intimal hyperplasia in a patient to an extentwhich makes it no longer necessary to surgically intervene, i.e., tore-establish a suitable blood flow through the vessel by repeatangioplasty, atheroectomy, or coronary artery bypass surgery. Theeffects of reducing, delaying, or eliminating stenosis may be determinedby methods routine to those skilled in the art including, but notlimited to, angiography, ultrasonic evaluation, fluoroscopic imaging,fiber optic endoscopic examination or biopsy and histology.

The amount of TGF-beta activator or production stimulator administeredis selected to treat vascular trauma of differing severity, with smallerdoses being sufficient to treat lesser vascular trauma such as in theprevention of vascular rejection following graft or transplant. TGF-betaactivators or production stimulators that are not characterized by anundesirable systemic toxicity profile at a prophylactic dose are alsoamenable to chronic use for prophylactic purposes with respect todisease states involving proliferation of vascular smooth muscle cellsover time (e.g., atherosclerosis, coronary heart disease, thrombosis,myocardial infarction, stroke, smooth muscle neoplasms such as leiomyomaand leiomyosarcoma of the bowel and uterus, uterine fibroid or fibromaand the like), preferably via systemic administration.

For prevention of restenosis, a series of spaced doses, optionally, insustained release dosage form, is preferably administered before andafter the traumatic procedure (e.g., angioplasty). The dose may also bedelivered locally, via catheter delivered to the afflicted vessel duringthe procedure. After the traumatic procedure is conducted, a series offollow-up doses are administered over time, preferably in a sustainedrelease dosage form, systemically to maintain an anti-proliferativeeffect for a time sufficient to substantially reduce the risk of or toprevent restenosis. A preferred therapeutic protocol duration afterangioplasty for this purpose is from about 3 to about 26 weeks.

High levels of lipoprotein Lp(a) are known to constitute a substantialrisk factor for atherosclerosis, coronary heart disease and stroke. Onesymptom associated with such conditions and other problems, such asrestenosis following balloon angioplasty and other pathogenicconditions, is the proliferation or the migration of smooth musclecells. No direct link between Lp(a) and proliferation of vascular smoothmuscle cells had been established in the prior art.

An in vivo pathway for the modulation of vascular smooth muscle cellproliferation is shown in FIG. 1. TGF-beta is believed to contribute tothe inhibitory mechanism that maintains vascular smooth muscle cells ina non-proliferative state in healthy vessels.

Vascular smooth muscle cell proliferation is inhibited by an active formof TGF-beta. Tamoxifen has been shown by the experimentation detailed inExample 1 hereof to stimulate both the production and the activation ofTGF-beta. Heparin stimulates the activation of TGF-beta by affecting therelease of the active form of TGF-beta from inactive complexes presentin serum. TGF-beta neutralizing antibodies inhibit the activity ofTGF-beta, thereby facilitating the proliferation of vascular smoothmuscle cells. An apparent in vivo physiological regulator of theactivation of TGF-beta is plasmin. Plasmin is derived from plasminogenthrough activation by, for example, TPA (tissue plasminogen activator).Plasmin activity is inhibited by the lipoprotein Lp(a) orapolipoprotein(a) (apo(a)), thereby decreasing the activation of thelatent form of TGF-beta and facilitating proliferation of vascularsmooth muscle cells.

An additional pathway for the modulation of vascular smooth muscle cellproliferation is shown in FIG. 2. Resting smooth muscle cells constitutecells in their normal, quiescent non-proliferative state. Such restingsmooth muscle cells may be converted to proliferating smooth musclecells through activation by platelet derived growth factor (PDGF),fibroblast growth factor (FGF) or other stimulatory moieties. Theproliferating smooth muscle cells may be converted to continualproliferating smooth muscle cells (ie., smooth muscle cells capable ofgenerating a pathological state resulting from over-proliferationthereof) by an autocrine growth factor. This growth factor is believedto be produced by proliferating smooth muscle cells. An increased levelof autocrine growth factor, which can be inhibited by the active form ofTGF-beta or an appropriately structured (ie designed) small moleculeinhibitor, is believed to mediate the production of continualproliferating smooth muscle cells.

Lp(a) consists of low density lipoprotein (LDL) and apo(a). Apo(a)shares approximately 80% amino acid identity with plasminogen (seeMacLean et al., Nature, 330: 132, 1987). Lp(a) has been found to inhibitcell-associated plasmin activity (see, for example, Harpel et al., Proc.Natl. Acad. Sci. USA, 86: 3847, 1989). Experiments conducted on humanaortic vascular smooth muscle cells derived from healthy transplantdonor tissue, cultured in Dulbecco's modified Eagles medium (DMEM)+10%fetal calf serum (FCS) as described in Kirschenlohr et al., Am. J.Physiol., 265, C571 (1993), indicated the following:

1) Addition of Lp(a) to sub-confluent human vascular smooth muscle cellsstimulated their proliferation in a dose dependent manner (addition of500 nM Lp(a) to human vascular smooth muscle cells caused a reduction indoubling time from 82+/−4 hours to 47+/−4 hours);

2) Addition of apo(a) had a similar effect, although a higherconcentration of apo(a) appeared to be required therefor;

3) Addition of LDL at varying concentrations up to 1 micromolar had noeffect on proliferation.

One possible mode of action for Lp(a) and apo(a) is competitiveinhibition of surface-associated plasminogen activation, which in turninhibits the subsequent activation of TGF-beta by plasmin. TGF-beta is apotent growth inhibitor of a number of anchorage-dependent cells,including smooth muscle cells. TGF-beta is produced as a latentpropeptide having a covalently linked homodimer structure in which theactive moiety is non-covalently linked to the amino-terminal portion ofthe propeptide. Latent TGF-beta must be cleaved (e.g., in vitro by acidtreatment or in vivo by the serine protease plasmin) in order to becomecapable of inhibiting the proliferation of vascular smooth muscle cells.Plasmin is therefore a leading candidate to be a physiological regulatorof TGF-beta.

The hypothesis that Lp(a) and apo(a) were acting on cultured humanvascular smooth muscle cells by interfering with activation of latentTGF-beta was tested. In support of this hypothesis, an observation wasmade that plasmin activity associated with vascular smooth muscle cellswas reduced 7-fold by Lp(a) and 5-fold by apo(a). The plasmin activityin the conditioned medium was also reduced by Lp(a) and apo(a) by about2-fold, but was much lower than cell-associated plasmin activity invascular smooth muscle cell cultures. These observations are consistentwith previous findings that Lp(a) is a more potent inhibitor ofsurface-associated, rather than fluid phase, plasminogen activation.

To exclude the possibility that Lp(a) was affecting the synthesis ofplasminogen activators rather than plasminogen activation, plasminogenactivator levels in human vascular smooth muscle cell cultures weremeasured in the presence and absence of the lipoproteins and in thepresence of a large excess of plasminogen, so that the lipoproteinspresent would not significantly act as competitive inhibitors. Totalplasminogen activator activity was not affected by the presence of anyof the lipoproteins in the vascular smooth muscle cell cultures. Forexample, plasminogen activator activity in the conditioned mediumremained at 0.7+/−0.6 mU/ml with Lp(a) additions up to 500 nM.

Lp(a) and apo(a) both reduced the level of active TGF-beta by more than100-fold compared to control or LDL-treated cultures. The level of totallatent plus active TGF-beta measured by ELISA as described in Example 8was unaffected by the presence of Lp(a) or apo(a), however. These factslead to the conclusion that Lp(a) stimulates proliferation of humanvascular smooth muscle cells by inhibiting plasmin activation of latentTGF-beta to active TGF-beta.

To further test this conclusion and exclude the possibility that Lp(a)was acting by binding active TGF-beta as well as reducing plasminactivity, human vascular smooth muscle cells were cultured in thepresence of Lp(a). These cells had a population doubling time of 47+/−3hours. Addition of plasmin was able to overcome the population doublingtime reducing effect of Lp(a) and reduce the cell number to controllevels, with the population doubling time increased to 97+/−4 hours.

The role of plasmin in the pathway was confirmed by studies in whichinhibitors of plasmin activity were added to human vascular smoothmuscle cells. Like Lp(a), these protease inhibitors increased cellnumber. Aprotinin, for example, decreased the population doubling timefrom 82+/−4 hours in control cultures to 48+/−5 hours, andalpha2-antiplasmin decreased the population doubling time to 45+/−2hours. 500 nM Lp(a) and aprotinin addition resulted in only a slightadditional stimulation of proliferation, with the population doublingtime for cultures of this experiment being 45+/−6 hours. Neutralizingantibodies to TGF-beta similarly decreased population doubling time invascular smooth muscle cells (see, for example, Example 1). In summary,Lp(a), plasmin inhibitors and neutralizing antibody to TGF-betastimulate proliferation of vascular smooth muscle cells, while plasminnullifies the growth stimulation of Lp(a). These results support thetheory that the mode of action of Lp(a) and apo(a) is the competitiveinhibition of plasminogen activation.

Experimentation conducted to ascertain the impact of tamoxifen onTGF-beta and vascular smooth muscle cell proliferation is set forth indetail in Example 1. The results of those experiments are summarizedbelow.

1) Addition of tamoxifen decreased the rate of proliferation, withmaximal inhibition observed at concentrations above 33 micromolar. 50micromolar tamoxifen concentrations produced a cell number 96 hoursfollowing the addition of serum that was reduced by 66%+/−5.2% (n=3) ascompared to cells similarly treated in the absence of tamoxifen.

2) Tamoxifen did not significantly reduce the proportion of cellscompleting the cell cycle and dividing. Inhibition of vascular smoothmuscle cells caused by tamoxifen therefore appears to be the result ofan increase in the cell cycle time of nearly all (>90%) of theproliferating cells.

3) Tamoxifen decreases the rate of proliferation of serum-stimulatedvascular smooth muscle cells by increasing the time taken to traversethe G₂ to M phase of the cell cycle.

4) Tamoxifen decreased the rate of proliferation of vascular smoothmuscle cells by inducing TGF-beta activity.

5) Vascular smooth muscle cells produced TGF-beta in response totamoxifen. Tamoxifen appears to increase TGF-beta activity in culturesof rat vascular smooth muscle cells by stimulating the production oflatent TGF-beta and increasing the proportion of the total TGF-betawhich has been activated.

6) Tamoxifen, unlike heparin, does not act by releasing TGF-beta frominactive complexes present in serum.

7) TGF-beta mRNA was increased by approximately 10-fold by 24 hoursafter addition of tamoxifen (10 micromolar). This result suggests thatthe expression of TGF-beta mRNA by the smooth muscle cells will beincreased, thereby facilitating decreased proliferation thereof byactivated TGF-beta.

8) Tamoxifen is a selective inhibitor of vascular smooth muscleproliferation with an ED₅₀ (a concentration resulting in 50% inhibition)at least 10-fold lower for vascular smooth muscle cells than foradventitial fibroblasts.

Additional experimentation has shown that the addition of Lp(a) orapo(a) substantially reduced the rat vascular smooth muscle cellproliferation inhibitory activity of tamoxifen, with the populationdoubling time in the presence of tamoxifen and Lp(a) being 42+/−2 hours(as compared to a population doubling time of 55+/−2 hours for tamoxifenalone, and a time of 35+/−2 hours for the control). Also, the presenceof Lp(a) reduced the levels of active TGF-beta produced in response tothe addition of tamoxifen by about 50-fold. Addition of plasmin to ratvascular smooth muscle cells treated with tamoxifen and Lp(a) resultedin most of the TGF-beta being activated, and proliferation was againslowed (with the population doubling time being 57+/−3 hours). Theseobservations are consistent with the theory that Lp(a) acts byinhibiting TGF-beta activation.

Identification of therapeutic agents (direct or indirect TGF-betaactivators or production stimulators) that act to inhibit vascularsmooth muscle cell proliferation by the pathway shown in FIG. 1 can beidentified by a practitioner in the art by conducting experiments of thetype described above and in Example 1. Such experimental protocolsfacilitate the identification of therapeutic agents useful in thepractice of the present invention and capable of one of the followingactivities:

1) production or activation of TGF-beta;

2) having TGF-beta-like activity;

3) activation of plasminogen;

4) increase in plasmin activity; or

5) reduction of Lp(a) or apo(a) level or levels of π-I or otherinhibitors of TGF-beta activation.

Identification of therapeutic agents (direct or indirect TGF-betaactivators or production stimulators) that act to inhibit vascularsmooth muscle cell proliferation by the pathway shown in FIG. 2 can beidentified by a practitioner in the art by conducting experimentationusing known techniques that are designed to identify growth factors madeby proliferating smooth muscle cells, which growth factors also act onthose cells (i.e., autocrine growth factors). Rational drug design canthen used to screen small molecules for the ability to inhibit theproduction or activity of such autocrine growth factors as leadcompounds for drug design. Such experimental protocols facilitate theidentification of therapeutic agents useful in the practice of thepresent invention and capable of one of the following activities:

1) production or activation of TGF-beta;

2) having TGF-beta-like activity; or

3) inhibit the activity or production of an autocrine growth factorproduced by proliferating smooth muscle cells.

Smooth muscle cell proliferation is a pathological factor in myocardialinfarctions, atherosclerosis, thrombosis, restenosis and the like.Therapeutic/prophylactic agents of the present invention, includingtamoxifen and the like, having at least one of the activities recitedabove and therefore being capable of inhibiting proliferation ofvascular smooth muscle cells, are useful in the prevention or treatmentof these conditions. Manipulation of the proliferation modulationpathway for vascular smooth muscle cells to prevent or reduce suchproliferation removes or reduces a major component of the arteriallesions of atherosclerosis and the restenosed arteries followingangioplasty, for example.

More specifically, chronically maintaining an elevated level ofactivated TGF-beta reduces the probability of atherosclerotic lesionsforming as a result of vascular smooth muscle cell proliferation.Consequently, administration of TGF-beta activators or TGF-betaproduction stimulators protects against atherosclerosis and subsequentmyocardial infarctions that are consequent to coronary artery blockage.Also, substantially increasing the activated TGF-beta level for a shorttime period allows a recipient to at least partially offset the strongstimulus for vascular smooth muscle cell proliferation caused by highlytraumatic injuries or procedures such as angioplasty. Continued deliveryto the traumatized site further protects against restenosis resultingfrom vascular smooth muscle cell proliferation in the traumatized area.

Prevention or treatment relating to a traumatized or diseased vascularsite, for example, the TGF-beta activators or production stimulators mayalso be administered in accordance with the present invention using aninfusion catheter, such as produced by C.R. Bard Inc., Billerica, Mass.,or that disclosed by Wolinsky (U.S. Pat. No. 4,824,436) or Spears (U.S.Pat. No. 4,512,762). In this case, a therapeutically/prophylacticallyeffective dosage of the TGF-beta activator or production stimulator willbe typically reached when the concentration thereof in the fluid spacebetween the balloons of the catheter is in the range of about 10⁻³ to10⁻² M. It is recognized by the present inventors that TGF-betaactivators or stimulators may only need to be delivered in ananti-proliferative therapeutic/prophylactic dosage sufficient to exposethe proximal (6 to 9) cell layers of the intimal or tunica media cellslining the lumen thereto. Also, such a dosage can be determinedempirically, e.g., by a) infusing vessels from suitable animal modelsystems and using immunohistochemical methods to detect the TGF-betaactivator or production stimulator and its effects; and b) conductingsuitable in vitro studies.

It will be recognized by those skilled in the art that desiredtherapeutically/prophylactically effective dosages of a TGF-betaactivator or production stimulator administered by a catheter inaccordance with the invention will be dependent on several factors,including, e.g.: a) the atmospheric pressure applied during infusion; b)the time over which the TGF-beta activator or production stimulatoradministered resides at the vascular site; c) the nature of thetherapeutic or prophylactic agent employed; and/or d) the nature of thevascular trauma and therapy desired. Those skilled practitioners trainedto deliver drugs at therapeutically or prophylactically effectivedosages (e.g., by monitoring drug levels and observing clinical effectsin patients) will determine the optimal dosage for an individual patientbased on experience and professional judgment. In a preferredembodiment, about 0.3 atm (i.e., 300 mm of Hg) to about 5 atm ofpressure applied for 15 seconds to 3 minutes directly to the vascularwall is adequate to achieve infiltration of a TGF-beta activator orproduction stimulator into the smooth muscle layers of a mammalianartery wall. Those skilled in the art will recognize that infiltrationof the TGF-beta activator or production stimulator into intimal layersof a diseased human vessel wall in free or sustained-release form willprobably be variable and will need to be determined on an individualbasis.

While two representative embodiments of the invention relate toprophylactic or therapeutic methods employing an oral dosage form orinfusion catheter administration, it will be recognized that othermethods for drug delivery or routes of administration may also beuseful, e.g., injection by the intravenous, intralymphatic, intrathecal,intraarterial, local delivery by implanted osmotic pumps or otherintracavity routes. Administration of TGF-beta activators or productionstimulators in accordance with the present invention may be continuousor intermittent, depending, for example, upon the recipient'sphysiological condition, whether the purpose of the administration istherapeutic or prophylactic and other factors known to skilledpractitioners.

In the practice of certain embodiments of the present invention,catheter administration routes including systemic and localized deliveryto the target site are preferably conducted using a TGF-beta activatoror production stimulator dispersed in a pharmaceutically acceptablecarrier. Tamoxifen and its structural analogs and salts, including thecompounds of formula (I) can be administered by a variety of routesincluding oral, rectal, transdermal, subcutaneous, intravenous,intramuscular, and intranasal. These compounds preferably are formulatedprior to administration, the selection of which will be decided by theattending physician. Typically, TMX and its structural analogs andsalts, including the compounds of formula (I), or a pharmaceuticallyacceptable salt thereof, is combined with a pharmaceutically acceptablecarrier, diluent or excipient to form a pharmaceutical formulation, orunit dosage form.

The total active ingredients in such formulations comprises from 0.1 to99.9% by weight of the formulation. By “pharmaceutically acceptable” itis meant the carrier, diluent, excipient, and/or salt must be compatiblewith the other ingredients of the formulation, and not deleterious tothe recipient thereof.

Pharmaceutical formulations containing TMX and its structural analogsand salts, including the compounds of formula (I), can be prepared byprocedures known in the art using well known and readily availableingredients. For example, the compounds of formula (I) can be formulatedwith common excipients, diluents, or carriers, and formed into tablets,capsules, suspensions, powders, and the like. Examples of excipients,diluents, and carriers that are suitable for such formulations includethe following fillers and extenders such as starch, sugars, mannitol,and silicic derivatives; binding agents such as carboxymethyl celluloseand other cellulose derivatives, alginates, gelatin, andpolyvinyl-pyrrolidone; moisturizing agents such as glycerol;disintegrating agents such as calcium carbonate and sodium bicarbonate;agents for retarding dissolution such as paraffin; resorptionaccelerators such as quaternary ammonium compounds; surface activeagents such as cetyl alcohol, glycerol monostearate; adsorptive carrierssuch as kaolin and bentonitc; and lubricants such as talc, calcium andmagnesium stearate, and solid polyethyl glycols.

The compounds also can be formulated as elixirs or solutions forconvenient oral administration or as solutions appropriate forparenteral administration, for example, by intramuscular, subcutaneousor intravenous routes.

The present invention also contemplates therapeutic methods andtherapeutic dosage forms involving sustained release of the TGF-betaactivator or production stimulator to target cells. Preferably, thetarget cells are vascular smooth muscle cells, cancer cells, somaticcells requiring modulation to ameliorate a disease state and cellsinvolved in immune system-mediated diseases that are accessible by localadministration of the dosage form. Consequently, the methods and dosageforms of this aspect of the present invention are useful for inhibitingvascular smooth muscle cells in a mammalian host, employing atherapeutic agent that inhibits the activity of the cell (e.g.,proliferation, formation of lipid proliferative lesions, contraction,migration or the like) but does not kill the cell and, optionally, avascular smooth muscle cell binding protein. Sustained released dosageforms for systemic administration as well as for local administrationare also employed in the practice of the present method. Formulationsintended for the controlled release of pharmaceutically-active compoundsin vivo include solid particles of the active ingredient that are coatedor tabletted with film-forming polymers, waxes, fats, silica, and thelike. These substances are intended to inhibit the dissolution,dispersion or absorption of the active ingredient in vivo.Hydroxypropylmethyl cellulose is one example of an ingredient that canprovide a slow or controlled release of the active ingredient. Thecompounds can also be delivered via patches for transdermal delivery,subcutaneous implants, infusion pumps or via release from implantedsustained release dosage forms.

Another embodiment of the invention relates to prophylactic ortherapeutic “sustained release” methods from the surface of anintravascular device employing an excipient matrix which will releasethe TGF-beta activators over a one-week to two-year or longer period.The surface coating and the impregnated forms of the article can be abiodegradable or nonbiodegradable polymer or ceramic material which willslowly release. the TGF-beta activator at a dose rate that will inhibitthe proliferation of fibromuscular cells and/or lipid accumulation whichwould impair the function of the device. The accumulation offibromuscular cells, including VSMC, and their associated matrix, alongwith lipid containing foam cells can decrease the lumenal area ofintravascular stents, synthetic grafts and indwelling catheters to anextent that blood flow is critically impaired and the device can failfunctionally. The inhibition of this proliferation would extend theclinically functional life of these devices and be of significantclinical benefit to the patients.

The sustained release dosage forms of this embodiment of the inventionneeds to deliver a sufficient anti-proliferative, preferably cytostatic,dosage to expose cells immediately adjacent to the device surface to betherapeutic. This would inhibit cellular attachment, migration andproliferation of the fibromuscular cells and foam cells. This dosage isdeterminable empirically by implanting a specific device intravascularlywith variable amounts of the TGF-beta activator and modification of thepolymer excipient, both of which would affect the rate and duration ofthe drug release required to achieve the cytostatic dosing which hasbeen demonstrated in vascular smooth muscle cell tissue cultureexperiments. Different types of devices may require different periods oftherapeutic drug release. For example, the use in grafts and stents areconsidered permanently implanted devices; however, it may not benecessary to have the active agent continuously released from thedevice. It appears from initial observations that if excessiveproliferation is prevented until the graft or stent is surrounded byquiescent tissue and covered by intact endothelium then continuedrelease of cytostatic agents may be unnecessary. Devices such asindwelling catheters, however, do not become embedded in quiescentvascular wall tissue and overgrown with endothelium. These devices mayrequire the continual release of drugs to suppress the proliferation oftissue over their external and lumenal surfaces. To achieve thisprolonged period of sustained drug release, larger amounts of agent anddifferent types of, or modification of, the polymer or excipient arepreferable.

The sustained release dosage forms of the present invention,particularly, for local administration, are preferably eithernon-degradable microparticulates or nanoparticulates or biodegradablemicroparticulates or nanoparticulates. More preferably, themicroparticles or nanoparticles are formed of a polymer containingmatrix that biodegrades by random, nonenzymatic, hydrolytic scissioning.A particularly preferred structure is formed of a mixture ofthermoplastic polyesters (e.g., polylactide or polyglycolide) or acopolymer of lactide and glycolide components. The lactide/glycolidestructure has the added advantage that biodegradation thereof formslactic acid and glycolic acid, both normal metabolic products ofmammals.

Therapeutic dosage forms (sustained release-type) of the presentinvention exhibit the capability to deliver therapeutic agent to targetcells over a sustained period of time. Such dosage forms are disclosedin co-pending U.S. patent application Ser. No. 08/241,844, filed May 12,1994, now abandoned which is a continuation-in-part of Ser. No.08/062,451, filed May 13, 1993, now abandoned which is in turn acontinuation-in-part of Ser. No. 08/011,669, now abandoned which is inturn a continuation-in-part of PCT application US 92/08,220, filed Sep.25, 1992. These applications are incorporated by reference herein.Therapeutic dosage forms of this aspect of the present invention may beof any configuration suitable for this purpose. Preferred sustainedrelease therapeutic dosage forms exhibit one or more of the followingcharacteristics:

microparticulate (e.g., from about 0.5 micrometers to about 100micrometers in diameter, with from about 0.5 to about 2 micrometers morepreferred) or nanoparticulate (e.g., from about 1.0 nanometer to about1000 nanometers in diameter, with from about 50 to about 250 nanometersmore preferred), free flowing powder structure;

biodegradable structure designed to biodegrade over a period of timebetween from about 3 to about 180 days, with from about 10 to about 21days more preferred, or nonbiodegradable structure to allow therapeuticagent diffusion to occur over a time period of between from about 3 toabout 180 days, with from about 10 to about 21 days preferred;

biocompatible with target tissue and the local physiological environmentinto which the dosage form is being administered, includingbiocompatible biodegradation products;

facilitate a stable and reproducible dispersion of therapeutic agenttherein, preferably to form a therapeutic agent-polymer matrix, withactive therapeutic agent release occurring through one or both of thefollowing routes: (1) diffusion of the therapeutic agent through thedosage form (when the therapeutic agent is soluble in the polymer orpolymer mixture forming the dosage form); or (2) release of thetherapeutic agent as the dosage form biodegrades; and

capability to bind with one or more cellular and/or interstitial matrixepitopes, with from about 1 to about 10,000 bindingprotein/peptide-dosage form bonds preferred and with a maximum of about1 binding peptide-dosage form per 150 square angstroms of particlesurface area more preferred. The total number bound depends upon theparticle size used. The binding proteins or peptides are capable ofcoupling to the particulate therapeutic dosage form through covalentligand sandwich or non-covalent modalities as set forth herein.

For example, nanoparticles containing a compound of the formula (I) maybe prepared using biodegradable polymers including poly(D,L-lacticacid)PLA, poly(D,L-lactic-co-glycolic) PLGA, methacrylic acid copolymer,poly(epsilon-caprolactone), using either 1) n-solventemulsification-evaporation techniques or 2) emulsification-precipitationtechniques. These processes involve dispersion of polymer in an organicsolvent (e.g., acetone or benzyl alcohol) with or without a co-solvent,typically methylene chloride. The compound of formula (I) is containedin the organic solvent. In some cases, solvents are then mixed and thenadded dropwise to an aqueous solution containing stabilizinghydrocolloid [e.g., poly(vinyl alcohol) or gelatin] (i.e., oil in water)with mechanical agitation or sonication. Following formation of thestable emulsion, the chlorinated solvent is removed via evaporation ofthe stirred emulsion, yielding nanoparticles that then can be freed oforganic solvents by tangential filtration or repeated washings bycentrifugation/resuspension. The resultant aqueous suspension can thenbe frozen with or without saccharide or other cryoprotectants andlyophilized to yield nanoparticles capable of resuspension inphysiological salt solutions with simple agitation or sonication.

Alternatively, the aqueous solution can be added with agitation orsonication to the organic phase lacking chlorinated solvent (i.e.,water-in-oil emulsion) followed by further addition of aqueous solutionto achieve a phase inversion, to precipitate the nanoparticles.Alternatively, precipitation can be augmented by addition to salting-outagents in the aqueous solvent. Typically, for emulsification-evaporationtechnique 750 mg PLGA can be dissolved in 30 mL of methylene chloride.Five mL of methylene chloride containing 75 mg of a compound of formula(I), for example, tamoxifen, is added. This organic phase is addeddropwise to 180 mL of aqueous solution of 2.5% poly(vinyl alcohol, PVP)(20-70 kD mol Wt.) with sonication using a Branson 450 sonifier at 15-55watt output, for approximately 10 minutes to form a soluble emulsion.Sonication is performed in an ice bath at a temperature not exceeding15° C. the emulsion is then further stirred at room temperature for 24hours to allow for evaporation of the chlorinated solvent. The resultantnanoparticles are purified further using a Sartorius targeted filtrationdevice fitted with a 100 mm pore polyolefin cartridge filter. For theemulsification-precipitation technique, 10 mL of aqueous PMP (10-30%w/w) is added, under mechanical stirring at 1200-5000 rpm, to 5 mL ofbenzyl alcohol containing 10-15% w/w polymer PLA or PLGA and 10-15 w/wof a compound of the formula (I), for example, tamoxifen, followingoil-in-water emulsion formation over 5 minutes. Water (160 mL) is thenadded to effect a phase inversion, resulting in diffusion of organicsolvent into the water with concomitant precipitation of polymer assolid nanoparticles in the ensuing 10 minutes.

For TGF-beta activators or production stimulators, such as compounds ofthe formula (I), several exemplary dosing regimens are contemplated,depending upon the condition being treated and the stage to which thecondition has progressed. For prophylactic purposes with respect toatherosclerosis, for example, a low chronic dose sufficient to, elevatein vivo TGF-beta production is contemplated. An exemplary dose of thistype is about 0.1 mg/kg/day (ranging between about 0.1 and about 10mg/kg/day), preferably about 0.1-1.0 mg/kg/day, most preferably about0.3 mg/kg/day. Another exemplary dose range is from about 0.01 to about1000 micrograms/ml. Such low doses are also contemplated for use withrespect to ameliorating stenosis following relatively low trauma injuryor intervention, such as vein grafts or transplants or organ allografts,for example.

For prevention of restenosis following angioplasty, an alternativedosing regimen is contemplated which involves a single “pre-loading”dose (or multiple, smaller pre-loading doses) given before or at thetime of the intervention, with a chronic smaller (follow up) dosedelivered daily for two to three weeks or longer following intervention.For example, a single pre-loading dose may be administered about 24hours prior to intervention, while multiple preloading doses may beadministered daily for several days prior to intervention.Alternatively, one or more pre-loading doses may be administered about1-4 weeks prior to intervention. These doses will be selected so as tomaximize TGF-beta activator or production stimulator activity, whileminimizing induction of synthesis and secretion of extracellular matrixproteins. Such a dosing regimen may involve a systemic pre-loading dosefollowed by a sustained release chronic dose, or the sustained releasedosage form may be designed to deliver a large dose over a short timeinterval as well as a smaller chronic dose for the desired time periodthereafter. Some nausea may be encountered at the higher dose; however,the use of a sustained release or other targeted dosage form is expectedto obviate this side effect, because the recipient will not be subjectedto a high systemic dose of the therapeutic agent.

The local particulate dosage form administration may also localize tonormal tissues that have been stimulated to proliferate, therebyreducing or eliminating such pathological (i.e., hyperactive)conditions. An example of this embodiment of the present inventioninvolves intraocular administration of a particulate dosage form coatedwith a binding protein or peptide that localizes to pericytes and smoothmuscle cells of neovascularizing tissue. Proliferation of thesepericytes causes degenerative eye disease. Preferred dosage forms of thepresent invention release compounds capable of suppressing thepathological proliferation of the target cell population. The preferreddosage forms can also release compounds that increase vessel lumen areaand blood flow, reducing the pathological alterations produced by thisreduced blood supply.

It will be recognized that where the TGF-beta activator or productionstimulator is to be delivered with an infusion catheter, the therapeuticdosage required to achieve the desired inhibitory activity can beanticipated through the use of in vitro studies. In a preferred aspect,the infusion catheter may be conveniently a double balloon or quadrupleballoon catheter with a permeable membrane. In one representativeembodiment, a therapeutically effective dosage of a TGF-beta activatoror production stimulator is useful in treating vascular trauma resultingfrom disease (e.g., atherosclerosis, aneurysm, or the like) or vascularsurgical procedures such as angioplasty, atheroectomy, placement of astent (e.g., in a vessel), thrombectomy, and grafting. Atheroectomy maybe performed, for example, by surgical excision, ultrasound or lasertreatment, or by high pressure fluid flow. Grafting may be, for example,vascular grafting using natural or synthetic materials or surgicalanastomosis of vessels such as, e.g., during organ grafting. Thoseskilled in the art will recognize that the appropriate therapeuticdosage for a given vascular surgical procedure (above) is determined inin vitro and in vivo animal model studies, and in human preclinicaltrials.

Sustained release dosage forms of an embodiment of the invention mayonly need to be delivered in an anti-proliferative therapeutic dosagesufficient to expose the proximal (6 to 9) cell layers of the tunicamedia smooth muscle cells lining the lumen to the dosage form. Thisdosage is determinable empirically, e.g., by a) infusing vessels fromsuitable animal model systems and using immunohistochemical, fluorescentor electron microscopy methods to detect the dosage form and itseffects; and b) conducting suitable in vitro studies.

In a representative example, this therapeutically effective dosage isachieved by determining in smooth muscle cell tissue culture thepericellular agent dosage, which at a continuous exposure results in atherapeutic effect between the toxic and minimal effective doses. Thistherapeutic level is obtained in vivo by determining the size, numberand therapeutic agent concentration and release rate required forparticulates infused between the smooth muscle cells of the artery wallto maintain this pericellular therapeutic dosage.

Human vascular smooth muscle cells (VSMC) are more difficult to grow inculture than VSMC derived from other species, such as rat. Mediumconditioned on human VSMC decreased the proliferation of rat VSMC invitro. Entry of rat VSMC into S phase of the cell cycle was notaffected. However, the duration of G₂ and/or M phase was extended.Anti-TGF-beta antibody reversed the delayed entry into M phase caused byexposure to human VSMC conditioned medium (HCM). An examination of theHCM showed that 64±12% of the TGF-beta present in the medium was alreadyactivated. In contrast, rat VSMC conditioned medium displayed very lowlevels of latent TGF-beta and no detectable TGF-beta activity. HumanVSMC were found to produce tissue plasminogen activator (TPA) activityin culture. The TPA leads to an increase in plasmin activity, which inturn activates TGF-beta. This was confirmed by culturing human VSMC inthe presence of aprotinin, a plasmin inhibitor. Aprotinin increased therate of proliferation of human VSMC to almost the same extent asneutralizing anti-TGF-beta antibodies and α₂-antiplasmin. Thus, growthof human VSMC in culture is determined by the production of TGF-betaactivated by plasmin, which feeds back in an autocrine loop to increasethe duration of the cell cycle.

Subcultured human aortic VSMC remain more differentiated in culture thanrat aorta VSMC (i.e., they contain higher levels of the smoothmuscle-specific isoforms of myosin heavy chain (SM-MHC) and α-actin).TGF-beta likely plays a role in maintaining SM-MHC and α-actin content,and thus may be responsible for maintaining cells in a moredifferentiated phenotype. In view of these data, heparin, which isbelieved to release TGF-beta from inactive complexes in the serum, wouldbe predicted to have little effect on the rate of proliferation of humanVSMC, which is already inhibited by endogenous active TGF-betaproduction. Such observations may explain why human clinical trials ofheparin administered after PTCA have failed to demonstrate anybeneficial effect.

Freshly dispersed rat aortic VSMC lose SM-MHC and α-SM actin as theystart to proliferate. After 7 days in culture when the cells reachconfluence, serum is removed, and approximately 40% of the VSMCreexpress SM-MHC and α-SM actin at levels comparable to those present infreshly dispersed cells. If the cells were subcultured for more thanfive passages and allowed to reach confluence, less than 1% reexpressSM-MHC even after prolonged serum withdrawal. These cells representproliferating de-differentiated VSMC.

When primary cultures of rat aortic VSMC are exposed to TGF-beta, theloss of the 204 kD (SM-1) and 200 kD (SM-2) SM-MHC isoforms issubstantially inhibited. However, TGF-beta did not induce re-expressionof SM-MHC in subcultured cells that have very low levels of thisprotein. Therefore, TGF-beta can maintain a cell's differentiated state(as defined by SM-MHC content), but cannot induce re-differentiation ina de-differentiated proliferating cell. Since TGF-beta extends the G₂phase of the cell cycle in both primary and passaged VSMC cultures, thedata suggest that the pathways that mediate proliferation anddifferentiation are regulated independently.

Specific markers of both differentiated and proliferating VSMCs havebeen isolated. Four cell populations were probed using generated cDNAs:(a) freshly dispersed rat aortic cells; (b) freshly dispersed rat aorticVSMC after 7 days in culture (D7 cells); (c) freshly dispersed rataortic VSMC after subculturing 12 times (S12 cells); and (d) ratfibroblasts. Five classes of gene markers were defined. Class 1 cDNAswere expressed to a similar level in all of the RNAs. Class 2 cDNAs werehighly expressed in RNA from freshly dispersed aortic cells, but werebarely detectable in D7 or S12 cells and were not detectable in ratfibroblasts. Class 3 cDNAs were expressed at similar levels in freshlydispersed aortic, D7 and S12 cells. Class 4 cDNAs showed higherexpression in freshly dispersed aortic and D7 cells than in S12 cellsand fibroblasts. Class 5 cDNAs were expressed more strongly in S12 cellsthan in freshly dispersed aortic cells, D7 cells and fibroblasts. Class4 genes included α-SM actin, γ-SM actin, SM22α, calponin, tropoelastin,phospholamban and CHIP28. In addition, previously defined markers of thedifferentiated phenotype include SM-MHC, integrin and vinculin. Class 5genes included matrix G1a (MGP) and osteopontin. When passaged cellswere made quiescent by removal of serum, the levels of MGP andosteopontin did not change significantly, indicating that highexpression of these two genes occurs in VSMC that have undergoneproliferation, but does not depend on the cells being in the cell cycle.

Such studies of gene expression provide insight into the processes ofde-differentiation that occur during proliferation of VSMC. In situhybridization analysis of balloon-injured rat carotid arteries suggeststhat dividing intimal cells present 7 days after injury express highlevels of both osteopontin and MGP RNA. In contrast, osteopontin is onlyweakly expressed in the media of intact rat aorta and carotid arteries.Osteopontin and MGP may play a role in regulating calcification, whichcan occur rapidly in vascular lesions.

In the course of investigating potential heterogeneity of cells from rataortas, three groups of VSMC clones have been identified. One groupconsists of small cells that have an epithelioid or cobblestonemorphology and proliferate without the need for added growth factors,suggesting production of an autocrine growth factor(s). The second groupconsists of intermediate size, spindle shaped cells that grow in acharacteristic “hills and valleys” pattern and are dependent onexogenous growth factors. These cells resemble the predominant cellmorphology in standard cultures of adult aortic VSMC. The third groupconsists of large, often multinucleate, cells with limited proliferativecapacity. These large cells express high quantities of smooth musclespecific proteins.

All three types of cells could be isolated from neonatal and adult rataortae. However, aortas from young rats yielded high proportions of thesmall cell clones, while those from adult rats yielded high proportionsof intermediate and large cell clones. Clones of small VSMC can beinduced to convert to intermediate sized cells by treatment withTGF-beta. A proportion of these cells, in turn, converts to large cellsif plated at low density. The small cells may represent a progenitorcell and the large, non-proliferating cells may represent mature VSMC.

VSMC derived from neonatal rat aortas differ from normal adult VSMC inseveral ways: (a) they do not require exogenous growth factors forsustained growth; (b) they secrete PDGF-like growth factors; (c) theygrow with a characteristic epithelioid morphology; and (d) they expresshigh levels of cytochrome P450IA1, elastin and osteopontin (J. Biol.Chem. 266:3981-86, 1991; Biochem. Biophys. Res. Comm. 177:867-73, 1991;Nature 311:669-71, 1984). After intimal damage, neointimal lesions growwith an epithelioid morphology, secrete a PDGF-like protein and displayincreased expression of osteopontin in the vascular wall (Proc. Natl.Acad. Sci. USA 83:7311-15, 1986). These data are consistent with thepresence in vivo of a subpopulation of VSMC that comprises a diminishingproportion of the total cell population with age and which proliferatespreferentially.

TGF-beta is released by platelets, macrophages and VSMC at sites ofvascular injury. Since VSMC and endothelial cells at the site ofvascular injury can synthesize and release t-PA, a local mechanism foractivating secreted TGF-beta exists. The level of t-PA activity dependson expression of plasminogen activator inhibitor-1 (PAI-1) which is alsosynthesized in the vessel wall, and may be up-regulated by TGF-beta. Inaddition, TGF-beta binds with high affinity to α2-macroglobulin. Suchbinding renders TGF-beta unable to bind to cell surface receptors forTGF-beta. Polyanionic glycosaminoglycans, such as heparin, are alsonormally present in the vessel wall, and these moieties can reverse theassociation of TGF-beta with α2-macroglobulin. The phenotypic state ofthe VSMC may affect the VSMC response to activated TGF-beta. Thephenotypic state of the VSMC may be influenced by their extracellularenvironment. Accordingly, the biological effects of TGF-beta are subjectto a variety of regulatory mechanisms.

TGF-beta inhibits DNA synthesis in rat aortic VSMC stimulated witheither PDGF or EGF. In serum stimulated cells, however, TGF-beta haslittle effect on DNA synthesis. Instead, TGF-beta exerts itsanti-proliferative effect by prolonging the G₂ phase of the cell cycle.Likewise, heparin inhibits proliferation of serum-stimulated rat VSMC byextending the G₂ phase of the cell cycle. This effect of heparin can beeliminated by anti-TGF-beta antibody. These observations suggest thatthe anti-proliferative effect of heparin on VSMC in vitro and possiblyin vivo may be exerted through the release of TGF-beta.

When VSMC are dispersed in cell culture, they lose contractile proteinsand modulate to a “synthetic” phenotype as they proliferate. Themajority of VSMC in atheromatous plaques appear to have this syntheticphenotype also. Since loss of smooth muscle-specific proteins occursspontaneously in cell culture in the absence of mitogens where noproliferation occurs, this phenotypic change is not attributable tomitogenic stimulation, but rather to removal of the cells from theirextracellular matrix. The matrix contains large quantities of collagenand glycosaminoglycans that may maintain VSMC in a contractile state.TGF-beta does not exert its anti-proliferative effect through inhibitionof phenotypic modulation, however, since it is effective at slowingproliferation of passaged cells that can no longer express contractileproteins. Thus, TGF-beta displays the independent properties of (1)maintaining differentiated adult VSMC in the contractile phenotype; (2)causing maturation of small VSMC to intermediate size, spindle-shapedVSMC; and (3) inhibiting VSMC proliferation regardless of phenotype.Change from a contractile to synthetic phenotype is not obligatory forproliferation.

Cultured VSMC synthesize and secrete large quantities of extracellularmatrix proteins. TGF-beta enhances production of extracellular matrixproteins, which favors maintenance of the synthetic phenotype in cellsthat have been allowed to modulate. In addition, TGF-beta increasesexpression of numerous protease inhibitors, which also increaseaccumulation of extracellular matrix proteins.

In hypertension, there is increased thickness of the vessel media, witha consequent decrease in maximum lumen diameter, leading to increasedvascular resistance. The increased thickness of the vessel media is dueto growth of VSMC within the media. In large conductance vessels, suchas the aorta, the VSMC growth is believed to be attributable primarilyto VSMC hypertrophy (i.e., enlargement of the cell withoutproliferation). In hypertensive animals, these vessels display anincreased incidence of polyploid cells within the aortic media. Inresistance vessels, such as the mesenteric arteries, however, VSMCproliferation may contribute to the increased thickness of the vesselmedia. Previously, VSMC growth in hypertension was believed to resultfrom elevated blood pressure. Current data suggest that increasedvascular tone and VSMC hypertrophy and/or hyperplasia may be causedindependently by a common stimulus. For instance, under certaincircumstances, the vasoconstrictor peptide AII may be mitogenic forVSMC. Further, VSMC stimulated with AII also synthesize TGF-beta. Thus,any mitogenic effect of AII might be inhibited by TGF-beta, with the neteffect of AII stimulation being arrest in G₁ and hypertrophy withoutproliferation. AII may induce activation of TGF-beta by stimulatingexpression of t-PA by VSMC.

The VSMC involved in hypertension remain within the media of the vesseland are surrounded by a heparin-containing extracellular matrix.Therefore, any TGF-beta produced is freely available and will maintainVSMC in a contractile state.

In obliterative vascular disease, such as atherosclerosis, VSMC migratefrom the media and proliferate in the intima. There they secreteextracellular matrix proteins and form a lipid-rich plaque thatencroaches on the vascular lumen. This process is similar to, but slowerthan, the process that occurs following PTCA, leading to restenosis.Such inappropriate intimal VSMC proliferation also occurs in vascularbypass grafts and the arteries of transplanted organs, leading to graftocclusion and organ failure, respectively. In atherosclerosis, the VSMCinvolved in the lesion are generally of the synthetic phenotype andlocalized in the intima, in contrast to the VSMC involved inhypertension.

For medial VSMC involved in atherosclerosis, VSMC migration isaccompanied by an increase in synthesis and secretion of matrix proteinsand by proliferation. TGF-beta may reduce or prevent the VSMCproliferative response to mitogens and/or may induce synthesis andsecretion of extracellular matrix proteins. The effect of TGF-beta inthis case would be reduction of cellularity and increase of the matrixcomponent of an atherosclerotic plaque.

Alternatively, VSMC in the intima may arise from a population ofneonatal-like VSMC that are capable of migration and preferentialproliferation following vascular injury. This intimal phenotype may beeither induced or selected in response to vessel injury. When thesecells are exposed to TGF-beta, the neonatal-like, small cell phenotypeshould convert into intermediate sized, spindle-shaped cells that nolonger produce an autocrine growth factor. Thus, cells of theintermediate size should have a decreased tendency to proliferate. Overtime, a portion of this intermediate sized population of cells wouldconvert to the large, non-proliferative VSMC phenotype.

If VSMC are producing autocrine TGF-beta, tamoxifen has minimal or nofurther inhibitory effect on VSMC proliferation. Moreover, theseTGF-beta-producing VSMC exhibit responses to mitogenic stimuli that maydiffer from those of VSMC that are not producing TGF-beta. Such dataprovides further evidence of a complex interaction between the elementsthat are likely involved in atherosclerosis and vascular injury ortrauma.

Transgenic mice that express the human apo(a) gene are useful tools forstudying TGF-beta activation, VSMC proliferation and vascular lesionsthat mimic early human atherosclerotic lesions. In these mice, theapo(a) accumulates in focal regions in the luminal surface of vesselwalls. These foci of apo(a) inhibit plasminogen activation, which leadsto a decrease in production of plasmin. A low local concentration ofplasmin results in reduced activation of TGF-beta. This inhibition ofTGF-beta activation is greatest at sites of highest apo(a) accumulation.Further, these effects are observed whether the transgenic mice are feda normal diet or a lipid-rich diet. Serum levels of activated TGF-betacorrelate with the immunofluorescence determinations performed on tissuesections. Osteopontin, a marker of activated VSMC, co-localized withfocal apo(a) accumulation and regions of very low TGF-beta activation.

The formation of the atherosclerotic lesion can occur in five stages:

1. MIGRATION. In a healthy vessel, most or all of the smooth musclecells (SMC) are contained in the vessel media. The appearance of SMC inthe enlarged intima during lesion formation must therefore requiremigration of the SMC from the media to the intima of the vessel.Inhibition of this SMC migration would significantly alter the nature ofthe lesion, and may ameliorate the pathology associated with lesionformation.

2. LIPID ACCUMULATION. Medial SMC in healthy vessel walls do notsignificantly accumulate lipid. However, intimal SMC have an increasedcapacity for lipid uptake and storage. When exposed to elevated levelsof circulating lipid (particularly low density lipoprotein; LDL), SMCmay become saturated with fatty lipid and die. The accumulation of lipidis necessary for the progression of the lesion to clinical significance,since it forms the thrombogenic necrotic core of the lesion. Inhibitionof lipid accumulation in the SMC should significantly reduce or preventlesion formation and/or progression, thus reducing or preventingatherosclerosis and resultant myocardial infarction.

3. RECRUITMENT OF INFLAMMATORY CELLS. Human lesions contain manymacrophage-derived cells. The process of recruitment, the function ofthese cells, and their contribution to pathology are unclear. Anoversimplified mechanism suggests that macrophages are attracted to thelipid accumulating in the lesion, in order to remove the lipid from thevessel wall. While inhibition of recruitment of macrophage-derived cellsmight reduce lesion pathology, it may also speed progression to thelipid-filled, rupture-prone state.

4. PROLIFERATION. Intimal SMC accumulation is accompanied by medialthinning in many cases. Therefore, total SMC number may not increasesignificantly at the lesion site. Furthermore, the chronic nature ofatherosclerosis makes it difficult to detect stimulation ofproliferation in these lesions. Data obtained from transgenic apo(a)mice suggest that apo(a) may stimulate SMC proliferation. However,evidence that SMC hyperplasia is the major contributor toatherosclerosis is lacking. Thus, the ultimate effect that inhibition ofapo(a) has on atherosclerosis is dependent on the contribution of SMCproliferation to initiation or progression of an atherosclerotic plaque.

5. EXTRACELLULAR MATRIX DEPOSITION. Atherosclerotic lesions are alsorich in extracellular matrix (ECM), and in particular, collagen fibers.Increased ECM synthesis may increase plaque stability. Early plaquerupture, leading to myocardial infarction, may be associated with lowECM deposition and resultant weakening of the fibrous cap that overlaysthe necrotic, lipid-rich core of the lesion.

Accordingly, atherosclerosis involves the complex interplay of variousprocesses, some of which may be yet unidentified. Targeting a singleprocess in an effort to reduce or prevent atherosclerosis depends onknowledge of the relative contribution of each process to the manifestedpathology. For these reasons, a coordinated, therapeutic strategy ispreferred. An exemplary strategy involves inhibition of SMC migration,lipid accumulation and proliferation, with possible beneficial effectsof increasing. ECM deposition.

A diagnostic assay for identifying patients at risk for atherosclerosis,and therefore for identifying suitable candidates for therapy, is alsoan embodiment of the invention. In addition, this diagnostic assayprovides a means to monitor patients that are being treated foratherosclerosis. In one format, a sandwich ELISA for determining totalTGF-beta, ELISA plates are coated with an antibody that binds bothlatent and active TGF-beta. Patient sera are incubated with these ELISAplates, then the plates are washed to remove unbound components of thepatients' sera. Rabbit anti-TGF-beta antibody, capable of binding bothlatent and active TGF-beta, is then added to the plates and incubated.The plates are then washed to remove unbound antibody, andperoxidase-labeled anti-rabbit IgG is added. After incubation andwashing, the plates are exposed to the chromogenic substrate,orthophenylenediamine. The presence of total TGF-beta in patients' serais then determined calorimetrically at A₄₉₂ by comparison to a standardcurve. In patients treated with an agent that modifies TGF-beta, apretreatment determination of TGF-beta can be compared withpost-treatment time points to monitor treatment results andeffectiveness.

In an alternate format, TGF-beta type II receptor extracellular domain,which recognizes the active form of TGF-beta, is coated onto ELISAplates. Patient sera are added to the plates, and processed as above.This assay measures active TGF-beta present in sera.

In another alternate format, fluorescent-labeled anti-TGF-beta antibodyor TGF-beta type II receptor extracellular domain is used in place ofperoxidase labeled second antibody to detect the presence of TGF-beta inpatients' sera. In yet another alternate format, anti-TGF-beta antibodyor TGF-beta type II receptor extracellular domain is labeled with aradioactive moiety capable of detection by standard means. These lattertwo assays may be performed in an ELISA format, with or without usingthe additional anti-TGF-beta antibody described above. In addition,these latter two assays are amenable to other automated or non-automatedassay and detection methods.

To determine whether an agent is a TGF-beta activator or TGF-betaproduction stimulator, an agent or mixture of agents is first tested onrat aortic vascular smooth muscle cells (rVSMCs) for their ability tostimulate the production of active TGF-β in the culture medium asoriginally described for tamoxifen. See Grainger et al. (Biochem. J.,294, 109 (1993)). The key step in demonstrating that cells have areduced proliferation rate as a result of TGF-β production andactivation is that the effect can be fully reversed by neutralizingantibodies to TGF-β. Incomplete reversal of a decreased rate ofproliferation is evidence for TGF-β independent effect(s), which mayinclude toxicity. The effects of an agent are then tested on explanthuman aortic smooth muscle cells (hVSMC) as described in Example 3 todetermine whether the agent also stimulates production of TGF-β by thesecells. The use of explant hVSMCs, prepared and grown as described inExample 3, is essential because (i) explant hVSMCs grown undernon-optimal conditions (particularly at low cell densities) willspontaneously produce TGF-β; (ii) hVSMC cultures from cells prepared byenzyme dispersal spontaneously produce substantial amounts of TGF-β inculture (Kirschenlohr et al., Am. J. Physiol., 265, C571 (1993)) andtherefore cannot be used for screening; and (iii) the sensitivity ofrVSMCs and hVSMCs to agents which induce the cells to produce TGF-βdiffers by up to 100-fold.

In screening for agents likely to be effective for clinical purposes, itis therefore necessary to use hVSMCs to determine both potency and thetherapeutic window between effective concentrations and toxicconcentrations for human cells. Candidate agents which pass the in vitrocell culture screens are then tested on one or more mouse models oflipid lesion formation. Efficacy of candidate agents is tested by theprotocols described in Example 7 for C57B 16 mice and mice expressingthe human apo(a) transgene that are fed a high fat diet, and also inapoE knockout mice fed a normal diet. Another animal model useful inscreening agents is the cholesterol-fed Watanabe rabbit. Finally, smallscale, pilot studies on candidate molecules are tested in patient groupswith clinically significant coronary artery disease for the ability ofthe drug to increase circulating concentrations of active TGF-β or toactivate latent forms of TGF-β.

The invention will be better understood by making reference to thefollowing specific examples.

EXAMPLE 1 Impact of Tamoxifen on Vascular Smooth Muscle Cells and theRelationship thereof to TGF-Beta Production and Activation

Cell Culture, DNA Synthesis Assay and Cell Counting

Rat vascular smooth muscle cells were cultured after enzymaticdispersion of the aortic media from 12-17 week old Wistar rats asdescribed in Grainger et al., Biochem. J., 277: 145-151, 1991. When thecells reached confluence (after about 6 days) the cells were releasedwith trypsin/EDTA (available from Gibco) and diluted 1:2 in Dulbecco'smodification of Eagle's medium (DMEM; available from ICN/Flow)supplemented with 100 U/mi penicillin and 10% fetal calf serum (FCS).The cells were then replated on tissue culture plastic (available fromICN/Flow) at approximately 1×10⁴ cells/cm². The cells were subculturedrepeatedly in this way when confluence was attained (about every 4days), and the cells were used between passages 6 and 12.

Rat adventitial fibroblasts were cultured as described in Grainger etal., Biochem. J., 283: 403-408, 1992. Briefly, the aortae were treatedwith collagenase (3 mg/ml) for 30 minutes at 37° C. The tunicaadventitia was stripped away from the media. The adventitia wasdispersed for 2 hours in elastase (1 mg/ml) and collagenase (3 mg/ml)dissolved in medium M199 (available from ICN/Flow). The cells were thenspun out (900×g, 3 minutes), resuspended in DMEM+10% FCS and plated outat 8×10⁴ cells/cm² on tissue culture plastic. When the cells reachedconfluence (after about 10 days), they were subcultured as described forvascular smooth muscle cells. Adventitial fibroblasts were subculturedevery 3 days at 1:3 dilution and used between passages 3 and 9.

DNA synthesis was assayed by [³H]-thymidine incorporation as describedin Grainger et al., Biochem. J., 277:145-151, 1991. Vascular smoothmuscle cells were subcultured, grown in DMEM+10% FCS for 24 hours, madequiescent in serum-free DMEM for 48 hours and restimulated with 10% FCSat “0” hours. [³H]-thymidine (5 microcuries/ml; available from AmershamInternational) was added 12 hours after restimulation and the cells wereharvested after 24 hours. DNA synthesis by adventitial fibroblasts wasdetermined similarly, except that the cells were made quiescent inserum-free DMEM for 24 hours.

Cells were prepared for counting by hemocytometer from triplicateculture dishes as described in Grainger et al., Biochem. J.,277:145-151, 1991. Cells were also counted by direct microscopicobservation of gridded culture dishes. The grids were scored into theplastic on the inner surface, so that the cells could not migrate intoor out of the area being counted during the experiment. Cells in each offour squares in two separate wells were counted at each time point. Allcell counting experiments were repeated on at least three separatecultures.

A stock solution of tamoxifen (5 mM; available from ICI Pharmaceuticals)was made up in 10% ethanol (EtOH) and diluted in DMEM and 10% FCS togive the final concentration. The effects of each tamoxifenconcentration were compared with the effects observed in control wellscontaining the same final concentration of the ethanol vehicle.Recombinant TGF-beta (available from Amersham International) wasdissolved in 25 mM Tris/Cl to give. a 5 microgram/ml stock solution andsterile filtered through a Spinnex Tube (such as a Centrex DisposableMicrofilter Unit available from Rainin Instrument Company, Inc., Woburn,Mass.). Neutralizing antiserum to TGF-beta (BDA19; available from R & DSystems) was reconstituted in sterile MilliQ water (available fromMillipore Corporation, Bedford, Mass.). At 10 micrograms/ml, thisantibody completely abolished the activity of 10 ng/ml recombinantTGF-beta on subcultured (8th passage) vascular smooth muscle cells.

Assays for TGF-Beta

The TGF-beta activity present in medium conditioned on various cells wasdetermined by DNA synthesis assay on mink lung endothelial (MvLu) cells;a modification of the assay described in Danielpour et al., J. Cell.Physiol., 138: 79-83, 1989. MvLu cells were subcultured at 1:5 dilutionin DMEM+10% FCS. After 24 hours, the medium was replaced with theconditioned medium to be tested in the absence or presence of theneutralizing antiserum to TGF-beta at 10 micrograms/mi. DNA synthesisduring a 1 hour pulse of [³H]-thymidine (5 microcuries/ml) wasdetermined 23 hours after addition of the test medium. TGF-beta activitywas calculated as the proportion of the inhibition of DNA synthesiswhich was reversed in the presence of neutralizing antibody, using astandard curve to convert the inhibition values into quantities ofTGF-beta. The TGF-beta standards and conditioned media both contained10% FCS in DMEM.

The total latent and active TGF-beta present was determined by asandwich ELISA (see Example 8). Maxisorb 96-well ELISA plates (availablefrom Gibco) were coated with neutralizing antiserum against TGF-beta(BDA19; available from R & D Systems) at 2 micrograms/cm² in phosphatebuffered saline (PBS) overnight at room temperature. The plates werewashed between each step with tris-buffered saline containing 0.1%Triton X-100 (available from Sigma Chemical Company). The plates wereincubated with samples for 2 hours, with a second antibody to TGF-beta(BDA5; available from R & D Systems) at 0.1 micrograms/ml for 2 hours,with anti-rabbit IgG. peroxidase- conjugated antibody (available fromSigma Chemical Co.) for 1 hour, and with the chromogenic substrateo-phenylenediamine (Sigma), made up according to manufacturer'sinstructions, for minutes. Absorbances at 492 nm were converted intoquantities of TGF-beta protein using a standard curve. Both conditionedmedia and standards were assayed in the presence of 10% FCS in DMEM.This assay was linear for TGF-beta concentrations in the range from 0.1ng/ml to 20 ng/ml in the presence of 10% FCS in DMEM.

RNA Preparation and Northern Analysis

Total cytoplasmic RNA was isolated from cultured vascular smooth musclecells as described in Kemp et al., Biochem. J., 277: 285-288, 1991.Northern analysis was performed by electrophoresis of total cytoplasmicRNA in 1.5% agarose gels in a buffer containing 2.2 M formaldehyde, 20mM 3-(N-morpholino)propanesulfonic acid, 1 mM EDTA, 5 mM sodium acetateand 0.5 micrograms/ml ethidium bromide. The integrity of the RNA waschecked by visualizing the, gel under UV illumination prior to transferonto Hybond N (available from Pharmacia LKB) as specified by themanufacturer. Filters were hybridized as described in Kemp et al.,Biochem. J., 277: 285-288, 1991, using a [³²P]-oligolabeled mouseTGF-beta probe corresponding to amino acids 68-228 in the precursorregion of the TGF-beta polypeptide as set forth in Millan et al.,Development, 11 1:131-144.

Results

Vascular smooth muscle cells from the aorta of adult rats proliferatewith a cell cycle time of approximately 35 hours in DMEM+10% FCS (see,for example, Grainger et al., Biochem. J., 277: 145-151, 1991). Additionof tamoxifen decreased the rate of proliferation with maximal inhibitionat concentrations above 33 micromolar. 50 micromolar tamoxifenconcentrations produced an increase in cell number (96 hours followingthe addition of serum) that was reduced by 66%+/−5.2% (n=3). The slowerrate of proliferation was hypothesized to stem from a complete blockageof proliferation for a proportion of the vascular smooth muscle cells orfrom an increase in the cell cycle time of all of the cells. Todistinguish between these possibilities, the proportion of the cellspassing through M phase and the time course of entry into cell divisionwere determined.

Quiescent vascular smooth muscle cells were stimulated with DMEM+10% FCSin the absence or presence of 33 micromolar tamoxifen, with the cellnumber being determined at 8 hour intervals by time lapsephotomicroscopy. In the presence of ethanol vehicle alone, more than 95%of the vascular smooth muscle cells had divided by 40 hours, whereasthere was no significant increase in cell number in the presence oftamoxifen until after 48 hours. By 64 hours, however, more than 90% ofthe cells had divided in the presence of tamoxifen. The time taken for50% of the cells to divide after stimulation by serum was increased from35 +/−3 hours (n=7) to 54+/−2 hours (n=3) by 33 micromolar tamoxifen.Since tamoxifen did not significantly reduce the proportion of cellscompleting the cell cycle and dividing, inhibition of vascular smoothmuscle cells caused by tamoxifen appears to be the result of an increasein the cell cycle time of nearly all (>90%) of the proliferating cells.

To determine whether tamoxifen increased the duration of the cell cycleof vascular smooth muscle cells by increasing the duration of the G₀ toS phase, the effect of tamoxifen on entry into DNA synthesis wasanalyzed. Tamoxifen at concentrations up to 50 micromolar did notsignificantly affect the time course or the proportion of cells enteringDNA synthesis following serum stimulation of quiescent vascular smoothmuscle cells (DNA synthesis between 12 hours and 24 hours afterstimulation was measured by [³H]-thymidine incorporation: control at17614+/−1714 cpm; 10 micromolar tamoxifen at 16898+/−3417 cpm; and 50micromolar tamoxifen at 18002+/−4167 cpm). Since the duration of S phaseis approximately 12 hours (unpublished data), tamoxifen does not appearto have significantly impacted the time course of entry into DNAsynthesis. These results therefore imply that tamoxifen decreases therate of proliferation of serum-stimulated vascular smooth muscle cellsby increasing the time taken to traverse the G₂ to M phase of the cellcycle.

Based upon these results, it appeared that tamoxifen exhibited effectssimilar to those previously described for TGF-beta (see, for example,Assoian et al., J. Cell. Biol. 109: 441-448, 1986) with respect toproliferation of subcultured vascular smooth muscle cells in thepresence of serum. Tamoxifen is known to induce TGF-beta activity incultures of breast carcinoma cell lines as described, for example, inKnabbe, et al., Cell, 48: 417-425, 1987. Consequently, experimentationwas conducted to determine whether tamoxifen decreased the rate ofproliferation of vascular smooth muscle cells by inducing TGF-betaactivity. When quiescent vascular smooth muscle cells were stimulatedwith 10% FCS in the presence of 50 micromolar tamoxifen and 10mincrograms/ml neutralizing antiserum against TGF-beta, the cellsproliferated at the same rate as control cells in the presence ofethanol vehicle alone.

To confirm that the vascular smooth muscle cells produced TGF-beta inresponse to tamoxifen, such cells were treated with tamoxifen for 96hours in the presence of 10% FCS. The conditioned medium was thencollected and TGF-beta activity was determined by the modified mink lungepithelial (MvLu) cell assay described above. Tamoxifen increased theTGF-beta activity in the medium by >50-fold. Addition of tamoxifen (50micromolar) in fresh DMEM+10% FCS to the MvLu cells had no effect on DNAsynthesis, demonstrating that tamoxifen did not induce production ofactive TGF-beta by the MvLu cells.

TGF-beta is produced as a latent propeptide which can be activatedoutside the cell by proteases such as plasmin. To determine whethertamoxifen increased TGF-beta activity by promoting the activation oflatent TGF-beta or by stimulating the production of the latentpropeptide which was subsequently activated, the total latent plusactive TGF-beta present in the conditioned medium was determined bysandwich ELISA as described above. After 96 hours in the presence oftamoxifen (50 micromolar), the total TGF-beta protein present wasincreased by approximately 4-fold. Furthermore, the proportion of theTGF-beta present in active form was increased from <5% in the mediumconditioned on vascular smooth muscle cells in the presence of ethanolvehicle alone to approximately 35% in the medium conditioned on cellstreated with tamoxifen. Thus, tamoxifen appears to increase TGF-betaactivity in cultures of rat vascular smooth muscle cells by stimulatingthe production of latent TGF-beta and increasing the proportion of thetotal TGF-beta which has been activated.

Heparin increases TGF-beta activity in medium conditioned on vascularsmooth muscle cells (unpublished data). The mechanism of action ofheparin in this regard appears to involve the release of TGF-beta frominactive complexes present in serum, because pretreatment of serum withheparin immobilized on agarose beads is as effective as direct additionof free heparin to the cells. To determine whether tamoxifen acts torelease TGF-beta from sequestered complexes in serum which are notimmunoreactive in the ELISA assay, 10% FCS+DMEM was treated with 50micromolar tamoxifen for 96 hours at 37° C. in the absence of cells.Medium treated in this way contained similar levels of TGF-beta proteinand activity to untreated medium. It appears, therefore, that tamoxifen,unlike heparin, does not act by releasing TGF-beta from inactivecomplexes present in serum.

The content of TGF-beta mRNA was also analyzed by Northern analysis atvarious time points after addition of tamoxifen. Subcultured ratvascular smooth muscle cells (6th passage in exponential growth) in theabsence or presence of ethanol vehicle alone contain very little mRNAfor TGF-beta. By 24 hours after addition of tamoxifen (10 micromolar),TGF-beta mRNA was increased approximately 10-fold.

Although TGF-beta decreases the rate of proliferation of vascular smoothmuscle cells, it does not affect the rate of proliferation offibroblasts. Tamoxifen at concentrations of up to 50 micromolar did notreduce the rate of proliferation of subcultured adventitial fibroblasts.Tamoxifen is therefore a selective inhibitor of vascular smooth muscleproliferation with an ED₅₀ at least 10-fold lower for vascular smoothmuscle cells than for adventitial fibroblasts.

EXAMPLE 2 Heparin Effect on VSMC Proliferation and Differentiation

Heparins

An unfractionated, high molecular weight, anticoagulant pig mucosalheparin, fragments of heparin devoid of anticoagulant activity, andfragments of heparin with anticoagulant activity were tested. Inaddition, heparin coupled to agarose beads (Sigma Chemical Co., St.Louis, Mo.) was examined (see also Grainger et al., Cardiovascular Res.27:223847, 1993).

Effect on Proliferation

Freshly dispersed rat VSMC, prepared as in Example 1, were cultured inmedium containing serum (as in Example 1) in the presence or absence ofheparin. The cells were counted at intervals. Depending on the heparinused, the increase in cell number at 144 hours (when control cells enterstationary phase) was reduced by between 27±4.2% and 76±3.2% (p<0.0005compared with cell number in control wells for all heparins tested).Although the effects of the heparins at 100 μg/ml were similar, therewas a trend to greater effectiveness with increasing molecular size. Thefour heparins of 20 kD or above inhibited proliferation by 60-76%, andthe four heparins of 12.6-3 kD inhibited proliferation by 27-45%.

Entry into Cell Cycle Phases

Heparin had no effect on the entry of cells into S phase, as determinedby growing the cells in the presence of 10 μM bromodeoxyuridine from0-72 hours. Similar results were obtained when the cells werepulse-labeled with [³H]-thymidine.

The proportion of cells completing mitosis in the presence or absence ofheparin was determined. Defined fields of cells were photographed ateight hour intervals by time lapse microscopy of gridded culture dishes.The grids were scored into the plastic on the inner surface so that thecells could not migrate into or out of the area being counted. In theabsence of heparin, 92±1% of primary cells divided by 60 hours, butthere was no detectable cell division in the presence of heparin until72 hours. By 88 hours, however, 96±2% of the cells had divided in thepresence of heparin. In the presence or absence of heparin, the time tocomplete mitosis was less than 3 hours. The total cell cycle times inthe presence and absence of heparin were determined. The data showedthat the major effect of heparin was to extend selectively the durationof G₂ to M phase of the cell cycle.

The concentration, of heparin required to inhibit S phase entrydecreased as the serum concentration was reduced. This observation isconsistent with the removal by heparin of components of serum requiredfor progression to S phase.

Heparin and TGF-beta

To determine whether TGF-beta mediated the effects of heparin,anti-TGF-beta antibody (10 μg/ml; R&D Systems) was added. Anti-TGF-betaantibody alone had no effect on VSMC proliferation stimulated by 10%FCS. This antibody completed reversed the inhibition of VSMCproliferation observed when cells were incubated in the presence ofheparin. Heparin coupled to agarose beads at an extracellularconcentration of 100 μg/ml was as effective as free heparin (100 μg/ml)at inhibiting VSMC proliferation. Agarose beads alone at the sameconcentration had no effect. These results are consistent withextracellular action of heparin on VSMC to inhibit proliferation.Further cell cycle studies indicated that heparin must be present withinthe first 12 hours of G₁ to inhibit VSMC proliferation.

Heparin and Smooth Muscle-specific Myosin Heavy Chain Expression

Previous studies demonstrated that primary VSMC in culture lose both the204 kD (SM-1) and the 200 kD (SM-2) isoforms of SM-MHC, whether the VSMCare cultured in serum or in serum-free medium onto fibronectin. Inprimary cultures stimulated by serum, 100 μg/ml heparin substantiallyinhibited the loss of both SM-1 and SM-2 proteins in all cells, asassayed by direct immunoperoxidase staining or Western blotting (CellTissues Res. 257:1137-39, 1989; Biochem. J. 277:145-51, 1991). If thecells were plated in serum-free medium onto fibronectin, the normal lossof SM-1 and MS-2 proteins was unaffected by the presence of heparin. Theeffect of heparin in preventing the de-differentiation of primary VSMCin serum was completely reversed by the addition of anti-TGF-betaantibody (10 μg/ml), indicating that this heparin effect was alsomediated by TGF-beta-like activity. Although heparin prevented the lossof smooth muscle-specific myosin heavy chain from primary VSMC in thepresence of serum, it did not promote its reexpression. Moreover,heparin did not promote reexpression of SM-MHC in subcultured cells thatexhibit very low levels of this protein. Thus, the effects of heparinand TGF-beta on the expression of SM-MHC in primary VSMC are similar.

EXAMPLE 3 Comparison of Enzyme-Dispersed and Explant-Derived Human VSMC

Materials

Collagenase (C-0130), elastase (E-0258), anti-rabbit IgGperoxidase-conjugated antibody, the chromogenic substrateorthophenylenediamine, and streptomycin sulfate were obtained fromSigma. Tamoxifen (free base) was purchased from Aldrich. Dulbecco'smodified Eagle's Medium (D-MEM) and medium M199 were purchased from FlowLaboratories. 6-[³H]-thymidine and the cell proliferation kit wereobtained from Amersham International. Anti-TGF-beta antibodies (BDA19and BDA47) were purchased from R&D Systems. EGF, PDGF-AA and PDGF-BBwere obtained from Bachem, and were dissolved in filter-sterilized 25 mMTris-HCl, pH 7.5, containing 1% fatty acid-free bovine serum albumin(BSA). Basic fibroblast growth factor and insulin-like growth factor 1(N-mer) were obtained from Bachem and dissolved in sterile MilliQ water.Antiotensin II and endothelin 1 were obtained from Sigma and dissolvedin sterile MilliQ water. TGF-beta (0.5 μg, lyophilized solid) waspurchased from Peninsula, dissolved in 5 mM HCl to yield a 5 μg/mlstock, and diluted with PBS+0.2% BSA.

Human Aortic VSMC Cultures

Adult human VSMC were obtained from 6 transplant donors (either sex, agerange from 3 to 54 years) using the enzyme dispersal or explanttechnique. In one case, the same donor (a 24 year old male) was used toestablish both an enzyme-dispersed (ED) and explant-derived (EX) cellculture. Prior to enzyme-dispersion or explanting treatment, humanaortas were obtained within 18 hours of death. The endothelium layer wasremoved with a scalpel blade and strips of smooth muscle cells (tunicamedia) were removed with forceps and chopped into small pieces (1 mm³).

ED Cultures

The aortic pieces were washed once with serum-free Hanks Balanced SaltSolution, then enzyme-dispersed with collagenase and elastase, asdescribed in Example 1. The cells were plated at an initial density of1.5×10⁵ cells/cm² and incubated in a humidified atmosphere at 37° C. in5% CO₂ in air. The cells were subcultured every 6-7 days (at stationaryphase) by releasing them with trypsin/EDTA and diluting them 1:1.5 inD-MEM+10% FCS. Subcultured ED cells were cultured with D-MEM+20% FCS 24h after plating, and thereafter at 48 hour intervals.

EX Cultures

The aortic pieces were washed once with D-MEM+10% FCS, resuspended in asmall volume of fresh D-MEM+10% FCS, and transferred to culture flasksor Petri dishes. The pieces were allowed to sediment onto the plasticand were evenly distributed (≈4 pieces/cm²). Cells started to grow outfrom the explants after 3-7 days in culture. The aortic pieces wereremoved during the third week in culture, and the cells adhering to theplastic were allowed to grow to confluence for a further week. The cellswere then subcultured every 4-5 days by releasing them with trypsin/EDTAand diluting them 1:2 in D-MEM+10% FCS. Subcultured cells were incubatedwith fresh D-MEM+20% FCS as described for ED cultures.

ED and EX subcultures were used between passage 5-20.

Cell counting, DNA synthesis assays and assays for total and activeTGF-beta were performed as described in Examples 1 and 8.

Results

ED and EX cultures prepared from the aorta of a single individualdisplayed distinct morphologies and growth characteristics. The EXculture proliferated much more rapidly than the ED culture. After 6weeks of subculturing the ED and EX culture whenever confluence wasattained, the total yield of cells was 4 fold higher per gram wet weightof aorta in the EX culture than the ED culture. The ED culture had, alonger population doubling time in D-MEM+20% FCS (71±5 hours) than theEX culture (35±2 hours).

The VSMC in the EX culture were spindle-shaped and grew to confluencewith a characteristic “hills and valleys” pattern at confluence. The EXculture VSMC reached stationary phase at a high saturation density(2.0-4.0×10⁴ cells/cm²). In contrast, the VSMC in the ED culture had astellate morphology with numerous long cytoplasmic projections. Theyreached stationary phase at a low saturation density (0.7-2.0×10⁴cells/cm²) without reaching monolayer coverage of the substrate. TheVSMC in the ED culture contained high levels of both SM-MHC and α-actin,while the VSMC in the EX culture contained much lower levels of both ofthese protein markers.

The longer population doubling time of human ED cultures compared to EDcultures from the rat aorta is due to autocrine production of activeTGF-beta. These human ED cultures produced 15.2±1.6 ng/ml total TGF-betaprotein, of which 64±12% was in the active form. In contrast, the humanEX cultures did not produce detectable amounts of TGF-beta. Mediumconditioned for 48 hours on EX cultures during exponential growthcontained <1 ng/ml total TGF-beta. When TGF-beta production was comparedusing ED and EX cultures obtained from the same donor, the ED cultureproduced 8.5 ng/ml total TGF-beta, of which 57% was in the active form.The corresponding EX culture produced <1 ng/ml total TGF-beta protein.

Exogenous TGF-beta (10 ng/ml) was added to EX cultures 24 hours aftersubculturing and cell number was determined at 24 hour intervals. After96 hours in the presence of exogenous TGF-beta, the increase in cellnumber was inhibited by 34±2%. The population doubling time of the EXcultures increased from 32±1 hour to 42±3 hours in the presence ofexogenous TGF-beta.

Because the addition of exogenous TGF-beta extended the populationdoubling time of EX cultures by less than 12 hours, TGF-beta activityalone cannot account for the difference in population doubling timebetween the ED and EX cultures. Therefore, the fraction of cells thatentered DNA synthesis in a 6 day period was compared usingbromodeoxyuridine incorporation with a cell proliferation kit. Theproportion of EX culture nuclei demonstrating bromodeoxyuridineincorporation after a 6 day pulse was 86±4%, but for ED culture cellswas 48±4%. Therefore, the population doubling time of ED cultures wasfurther increased over that of EX cultures, because less of the ED cellsthan the EX cells were cycling in the presence of D-MEM+20% FCS.

Tamoxifen (TMX) inhibits proliferation of rat ED VSMC by inducingTGF-beta production with a half-maximal inhibition of proliferation at2-5 μM TMX. Because human ED cultures already produce autocrineTGF-beta, the addition of TMX would not be expected to reduce the rateof VSMC proliferation further. To confirm this prediction, variousconcentrations of TMX (1 nM to 100 μM) or ethanol vehicle only (20 ppmto 0.2%) were added to the human VSMC for 96 hours, and the cell numberwas determined by cell counting. Concentrations of TMX >33 μM causedcell death, but concentrations below 10 μM did not affect the rate ofproliferation.

EX cultures of human VSMC did not produce autocrine TGF-beta, so TMXwould be predicted to inhibit VSMC proliferation. Concentrations of >33μM TMX caused cell death in human EX cultures, as observed with human EDcultures. The half-maximal inhibitory dose for EX cultures was 30-100 nMTMX. At 5 μM TMX, the increase in cell number in human EX cultures wasinhibited 33±8%.

To confirm these observations, quiescent EX cultures were restimulatedand cultured for 96 hours in D-MEM+20% FCS containing TMX (0.5 μM) inthe presence or absence of anti-TGF-beta antibody (25 μg/ml). Theincrease in cell number in the presence of TMX alone was inhibited by27±2%, as compared to control cells incubated with ethanol vehiclealone. The presence of anti-TGF-beta antibody completely reversed theinhibition of proliferation due to TMX. ELISA assays for TGF-betaconfirmed that medium conditioned on human EX cultures in the presenceof 5 μM TMX contained 6.0±2.0 ng/ml total TGF-beta protein, of which55±5% was activated.

The effect of heparin on proliferation of human ED and EX cultures wasexamined. Heparin IC86-1771, known to inhibit proliferation of rat EDVSMC by releasing a TGF-beta-like activity from serum, partiallyinhibited the proliferation of human EX cultures, but not ED cultures.At 100 μg/ml and at 48 hours after addition, heparin inhibited theincrease in cell number in EX cultures by 51±10%; at 96 hours afteraddition, by 71±15%. In ED cultures at 96 hours after addition of 100μg/ml heparin, the increase in cell number was inhibited by 8±5%.Anti-TGF-beta antibody did not abolish the ability of heparin to inhibitthe proliferation of human EX cultured VSMC. Therefore, human EX VSMCmay release more TGF-beta from 20% FCS than could be neutralized byadded antibody, or heparin affected TGF-beta DNA synthesis as well asTGF-beta activation at the heparin concentrations tested.

The effect of mitogens on the entry of ED and EX cells into DNAsynthesis was examined. Quiescent ED and EX VSMC were restimulated witheither 20% FCS or 100 ng/ml PDGF-BB in D-MEM, and entry into DNAsynthesis was monitored during successive 8 hour pulses using[³H]thymidine. EX cells entered DNA synthesis in response to bothmitogenic stimuli more rapidly than ED cells. The EX cells reached peakrate of DNA synthesis in response to FCS 16-24 hours after stimulation.The ED cells reached peak rate of DNA synthesis 24-32 hours aftermitogenic stimulation.

Quiescent EX cells were then exposed to various mitogens, andstimulation of DNA synthesis was determined by incorporation of[³H]thymidine 16-32 hours after stimulation. DNA synthesis wasstimulated by 20% FCS by 8.0±1.5 fold, compared to control cells thatremained in serum-free D-MEM throughout. PDGF-BB and PDGF-AA caused a≈3.0 fold stimulation of DNA synthesis. Insulin-like growth factor(IGF-1; 25 ng/ml) provided a 1.2 fold stimulation. However, epidermalgrowth factor (EGF; 100 ng/ml), basic fibroblast growth factor (bFGF;100 ng/ml), TGF-beta (10 ng/ml), angiotensin II (AII; 100 nM) andendothelin-1 (ET-1; 100 nM) did not significantly stimulate DNAsynthesis.

Quiescent ED cells were exposed to various mitogens, and stimulation ofDNA synthesis was determined by [³H]thymidine incorporation 16-40 hoursafter stimulation. DNA synthesis was stimulated by 20% FCS by 25±6 fold,compared to control cells that remained in serum-free D-MEM throughout.PDGF-BB stimulated ≈3.0 fold, but PDGF-AA stimulated only 2.0 fold. Thelatter response was also variable (1 of 3 cultures did not respond toPDGF-AA), in contrast to the stimulation of EX VSMC. IGF-1 and EGFstimulated DNA synthesis 1.3 fold, and bFGF, TGF-beta, AII. and ET-1 didnot stimulate DNA synthesis.

EXAMPLE 4 TGF-beta and Transgenic apo(a) Mice

Apo(a) mice

Human apo(a) has been expressed in transgenic mice (Nature360:670-72,1992), a species that normally lacks apo(a). These mice wereused to study whether inhibition of TGF-beta activation, resulting inenhanced VSMC proliferation, represents a key step in atherogenesis.

Apo(a) transgenic mice, when fed a lipid-rich diet, develop vascularlesions similar to the fatty streak lesions in early humanatherosclerosis. Immunoperoxidase labeling showed that apo(a)accumulated in the vessel wall at strongly staining focal regions in theluminal surface of the vessel. This phenomenon was studied using themore sensitive technique of immunofluorescence labeling.

Briefly, transgenic apo(a) mice, confirmed for the presence of theapo(a) gene by Southern blotting, and normal litter mates were obtainedby continued crossing of transgenic mice with C57/B16×SJL hybrids. Theheart and attached aorta were dissected out, immediately frozen inliquid nitrogen, embedded, and 6 μm frozen sections were prepared. Thesections were fixed in ice-cold acetone for 90 seconds and stored at−20° C. until used. All fluorescent labeling procedures were performedat 4° C. For apo(a) immunolabeling, sections were incubated with 3% BSAin Tris-buffered saline (TBS) for 30 minutes, then with sheep anti-humanLp(a) antibody that had been adsorbed against human plasminogen diluted1:1000 in TBS containing 3% BSA. The anti-human Lp(a) antibody had nodetectable cross-reactivity with mouse plasminogen. The bound primaryantibody was detected using fluorescein-conjugated rabbit anti-sheep IgGdiluted 1:80 in TBS containing 3% BSA, and visualized by fluorescencemicroscopy at 400×magnification (λexc=440nm; λem=510 nm);photomicrographs were taken with 5 second exposures (ASA 1600). Thetissue sections were indistinguishable whether the mice were fed anormal diet (Techlad, Madison, Wis.; 4% mouse/rat chow) or a lipid-richdiet containing 1.25% cholesterol, 7.5% saturated fat as cocoa butter,7.5% casein and 0.5% sodium chelate.

Immunofluorescence labeling for apo(a) showed strongly labeled foci ofapo(a) in the luminal surface of the aortic wall, but apo(a) was alsolabeled at a substantially lower intensity throughout the media of thevessel. No apo(a) labeling was detected in the aortic sections from thenormal litter mate mice. The serum concentration of apo(a) in thetransgenic mice was 3.8±1.2 mg/dl. Analysis of human arteries and ofmice injected with radiolabeled apo(a) showed that plasma-derived apo(a)penetrates the vessel wall. In situ hybridization suggested that little,if any, apo(a) in the vessel wall of the apo(a) mice was derived fromlocal synthesis.

Total and Activated Plasminogen

Activation of plasminogen in the aortic wall was assayed using thespecific inhibitor, α2-antiplasmin ((α2-AP), which forms a stablecovalent conjugate with active plasmin, but does not bind covalently toplasminogen, apo(a) or other proteins in the vessel wall. Briefly, α2-AP(Sigma) was labeled with either fluorescein isothiocyanate (Sigma) ortrimethylrhodamine isothiocyanate (Experimentia 16:430, 1960), andseparated from unincorporated label by two gel filtrations on SephadexG25.

For determination of activated plasminogen, sections were incubated for16 hours with α2-AP-FITC (1 μg/ml) and washed. For determination oftotal plasminogen, the sections were incubated with (α2-AP-FITC, asabove, washed thoroughly in TBS containing 0.2% Nonidet-P40 (NP-40) and300 mM NaCl (wash buffer), and then incubated with 1 mg/ml recombinanthuman tissue plasminogen activator (rTPA) in TBS for 3 hours to activatethe plasminogen. The sections were washed, incubated for 16 hours withα2-AP-TRITC (1 μg/ml), then washed thoroughly in wash buffer, followedby TBS. Bound labeled α2-AP was visualized by fluorescence microscopy at400×magnification (λexc=440 nm; λem=510 nm for FITC label; λexc=490 nm;λem=580 nm for TRITC label). The low level of backgroundautofluorescence from the acetone-fixed sections was subtracted for eachsection from the fluorescence of the label. There were no significantdifferences in the autofluorescence intensity either between sectionsfrom the same mouse aorta, or between normal litter mate aortic sectionsand those from transgenic apo(a) mice. Photomicrographs of boundα2-AP-FITC to detect active plasmin were exposed for 10 seconds, and ofbound α2-AP-TRITC to detect plasminogen were exposed for 1 second (1600ASA).

Quantitation of Fluorescence

A Magiscan image analysis system (Joyce-Loebl) with extended linearrange TV camera (Photonic Science) attached to a Nikon Diaphor invertedfluorescence microscope was used to quantitate the fluorescence. Thegain control on the photomultiplier was set so that the average pixelvalue over the area of the vessel wall was between 2-5% of full scale.For each section, four fields of aortic wall were selected randomlyunder phase contrast (400×magnification), and separate fluorescenceimages were captured using filters for fluorescein andtrimethylrhodamine. For TGF-beta and plasminogen/plasmin, the averagepixel value for the fluorescence intensity over the whole area of thevessel media was calculated, and the mean for the four sections fromeach mouse (i.e., 16 fields of view) was computed. For osteopontin, thevessel media was only partly labeled, and only pixels with intensityvalues >5% of full scale were included in the calculation of averagepixel value. The number of pixels (×10⁻²) above the threshold is shownas the area labeled for osteopontin.

The α2-AP-FITC was detected in aortic sections of both the normal andapo(a) mice, predominantly associated with the elastic laminae of thevessels. Quantitation of the fluorescent label showed approximately 3fold less active plasmin in the vessel wall of the apo(a) mice than inthe normal mice, regardless of whether the mice had been fed alipid-rich or normal diet, as shown in Table 1.

TABLE 1 Quantitative fluorescent data Normal Mice Transgenic apo(a) MiceNormal Diet Lipid-Rich Normal Diet Lipid-Rich TGF-β Total 112 ± 7  95 ±12 115 ± 1  109 ± 6  % Active 90 ± 6  90 ± 5  36 ± 3* 46 ± 8*Plasminogen Total 702 ± 47  748 ± 95  789 ± 121 688 ± 133 % Active 6.3 ±1.3 6.1 ± 0.6  1.7 ± 0.7*  1.9 ± 1.2* Osteopontin Total 1.4 ± 0.8 0.4 ±0.1 32.3 ± 4.4*   12.6 ± 2.1*⁺ Area 0.7 ± 0.9 1.2 ± 1.6 80.3 ± 0.0*   103 ± 31.7*⁺ *p < 0.05 for apo(a) mice compared with normal littermate mice^(+p < 0.05 for apo(a) mice on a lipid-rich diet compared with apo(a) mice on a normal diet (Student's unpaired t-test))

Control experiments demonstrated that the α2-AP-FITC bound only toactive plasmin in the sections. No fluorescence was detected in aorticsections that were incubated with α2-AP-FITC in the presence of a largeexcess (1 mU) of exogenous active plasmin. Aortic sections were alsoincubated with α2-AP-FITC after treatment with the plasmin inhibitor,aprotinin (100 μg/ml), and no fluorescence was detected, demonstratingthat there was no interaction of the label with the sections in theabsence of active plasmin.

To assay for plasminogen, active plasmin was first labeled withα2-AP-FITC, as described above, then the same sections were treated withrTPA to activate the plasminogen. The sections were relabeled for activeplasminogen using α2-AP-TRITC. When the rt-PA was omitted, no furtherstaining for active plasmin with α2-AP-TRITC was observed. Quantitationof the two fluorescent labels of active plasmin before and afteractivation of the plasminogen provides a measure of the total amount ofplasminogen and of the proportion of plasminogen that was alreadyactivated in the sections (see Table 1). There was no significantdifference in the total amounts of plasminogen in the sections from theapo(a) mice and normal mice. In the normal mice, ≈6% of the plasminogenwas activated to plasmin, compared with only 2% in the apo(a) transgenicmice. Thus, apo(a) inhibits plasminogen activation.

TGF-beta

To determine whether the low plasmin concentration in the aortic wall ofthe apo(a) mice resulted in reduced activation of TGF-beta,immunofluorescent labels were used to quantitate active TGF-beta andtotal TGF-beta (active+latent). Briefly, sections prepared as describedabove were labeled for total TGF-beta for 2 hours with 25 μg/ml of BDA47(R&D Systems), a rabbit polyclonal antiserum to TGF-beta that detectsisoforms 1 and 3 with equal sensitivity, but does not distinguishbetween latent and active TGF-beta. The sections were washed 3 times inTBS, and incubated with goat anti-rabbit IgG (Sigma; 1:50 dilution)conjugated with TRITC. Both antibodies were diluted in TBS containing 3%BSA. The same section was then washed 3 times in TBS and labeled foractive TGF-beta with R²X (TGF-beta type II receptor extracellulardomain, which recognizes the active form of isoforms 1 and 3 only) thatwas conjugated with FITC, as described above. Sections were incubatedfor 16 hours, then washed 3 times in PBS. Bound label was visualized byfluorescence microscopy, as described above. Photomicrograph exposureswere 5 seconds (1600 ASA). To calibrate the fluorescence intensities ofthe two labels, a solution containing various proportions of activeTGF-beta (6 ng/ml of total TGF-beta) was spotted ongelatin-polylysine-coated slides and allowed to dry at room temperature.The protein spots were labeled for total and active TGF-beta, asdescribed for the aortic sections, and the fluorescence intensity ratios(TRITC/FITC) were determined. False color images of the proportion ofTGF-beta in the active form were computed from the fluorescence ratiosof the aortic sections using the calibration.

TGF-beta was present throughout the aortic media, predominantlyassociated with the elastic laminae in both the normal and apo(a) mice.No fluorescent label was bound to the sections when the primaryanti-TGF-beta antibody was omitted. Quantitation of the fluorescentlabel showed no significant difference in the total amount of TGF-betapresent in the aortic wall of normal and apo(a) mice (see Table 1).

Active TGF-beta was assayed using a truncated extracellular domain ofthe type II TGF-beta receptor fused to glutathione-S-transferase (R2X)that had been FITC labeled. This label was detected in sections fromboth normal and apo(a) mice in association with the elastic laminae. Inthe presence of 100 mg/mi recombinant active TGF-beta-1, the binding ofR2X-FITC to the sections was completely blocked. In addition,glutathione-S-transferase labeled with FITC did not detectably bind toaortic sections from either normal or apo(a) mice.

The TGF-beta present in the aortic wall from apo(a) mice wassignificantly less active than the TGF-beta in the aortic wall fromnormal mice, irrespective of whether the mice had been fed a lipid-richdiet or normal diet (see Table 1). Thus, TGF-beta activation in theaortic wall is significantly inhibited by the presence of apo(a).Moreover, activation of TGF-beta is most strongly inhibited at the sitesof highest apo(a) accumulation. Therefore, changes in the vessel wallthat are a consequence of reduced TGF-beta activity will occurpreferentially at the sites of focal apo(a) accumulation, but will notbe dependent on the accumulation of lipid.

The mouse serum was also assayed for inhibition of TGF-beta activationby apo(a), using ELISAs for total and active TGF-beta (see Example 8).The total TGF-beta in the serum of apo(a) mice was 14.4±4.7 ng/ml; innormal mice it was 14.2±3.5 ng/ml. However, the proportion of totalTGF-beta that was active in the serum of apo(a) mice was 34±19%,compared with 92±12% active TGF-beta in the serum of normal mice.

Osteopontin

Aortic sections were assayed for osteopontin, a marker of activatedsmooth muscle cells. Osteopontin was detected by incubating sectionswith monoclonal antibody MPIIIB 10₁ (National Institute of HealthDevelopmental Studies Hybridoma Bank) at 10 μg/ml in TBS containing 3%BSA for 16 hours. The sections were washed 3 times in TBS, and boundantibody was detected using goat anti-mouse IgG conjugated tofluorescein (Sigma F-2012; 1:50 dilution; 2 hours). Photomicrographswere obtained with 2.5 sec exposure time (ASA 1600).

Fluorescent labeling of osteopontin was detected in the aortic sectionsfrom apo(a) mice on either a lipid-rich or normal diet. Although a smallincrease in labeling for osteopontin was detected throughout the mediaof the aortae from transgenic apo(a) mice, very high levels ofosteopontin labeling were co-localized with regions of focal apo(a)accumulation and very low TGF-beta activation. Treatment of apo(a) micewith bromodeoxyuridine for 24 hours before sacrifice showed nosignificant mitotic activity in the aortic media. Thus, in the absenceof physical injury, replication rates in atheromatous plaques are low,reflecting the slow growth of the lesions. Areas of aortic sections fromnormal mice that showed high proportions of active TGF-beta did not showdetectable labeling for osteopontin. The total intensity and area ofosteopontin labeling in the normal mouse sections were also very lowcompared with the apo(a) mouse sections. Therefore, the presence ofapo(a) induces osteopontin expression in VSMC in the aortic wall,similar to the changes that occur during the development of vascularlesions, regardless of whether the mice are fed a lipid-rich or normaldiet. Accumulation of lipid into the vessel wall under conditions wherecirculating lipid is elevated may be a consequence, rather than a cause,of the changes in VSMC activation marked by the expression ofosteopontin. Previous studies have shown that activated VSMC in cultureaccumulate about 20 fold more lipid than contractile VSMC.

The results of these experiments link apo(a) to the inhibition ofplasminogen and latent TGF-beta activation. The inhibition of TGF-betaactivation likely contributes to the subsequent development of fattylesions when apo(a) containing subjects (mice or human) are subject to alipid-rich diet.

EXAMPLE 5 Tamoxifen Inhibits Migration and Lipid Uptake in VSMC in vitroand in Transgenic Mice

Cell Culture

Rat aortic VSMCs from 12-20 week old Wistar male rats were prepared byenzyme dispersion, as described in Example 1. The cultured cells wereconfirmed as >99% SMC by staining for SM-MHC, and proliferated with acell cycle time of 36 h. Cells were passaged as described in Example 1,and were used either in primary culture or between passages 6-12.

Human aortic SMC from donors of either sex, aged 15-60, were prepared byexplanting 1 mm³ of medial tissue, as described in Example 3.

Migration

Migration was assayed using SMC grown to confluence on glass coverslips.A defined injury is performed on the confluent layer of cells, which areallowed to recover in D-MEM+10% FCS for 24 hours. Bromodeoxyuridine (10μM) is added between 18-24 hours, to label proliferating cells. Cellsmigrating past the boundary of the wound edge at 24 hours are detectedby propidium iodide (PI) staining of the cell nuclei (500 μM PI inPBS+0.5% NP-40 for 30 min at room temperature). Cells that synthesizedDNA were detected by antibody staining for bromodeoxyuridine usingfluorescein-conjugated anti-bromodeoxyuridine antibodies. Migrating andproliferating cells in each field of view were simultaneously counted byimage analysis of the rhodamine emission from PI and fluoresceinemission from bromodeoxyuridine.

Lipid Uptake

Cells in 24 well plastic dishes were incubated with serum-free D-MEM for24 hours or 1 hour at 37° C., then washed in PBS+1% BSA at 4° C. on icefor 30 minutes. Cells were incubated with ¹²⁵I-labeled LDL at variousconcentrations for 3 hours in the presence or absence of cold competitorLDL. The cells were washed six times with ice-cold PBS, lysed in 0.1 MNaOH or 0.1% SDS, and cell-associated counts of LDL were determined bygamma counting.

Apo(a) Transgenic Mice

Apo(a) [human 500 kD isoform] was expressed from the transferrinpromotor in C57/B16×SJL F1 cross mice. Mice were sacrificed at 24 weeksof age after 12 weeks on a lipid-rich or normal diet. Heart/lung/aortaefrozen blocks were prepared, and 6 μum frozen sections prepared ongelatin-coated slides. Sections were either fixed in acetone for 90seconds (for quantitative immunofluorescence; QIF) or in formaldehydevapor for 18 hours (for histology). Sections were stored at −20° C.until analyzed.

Histology

Sections were stained with trichrome stain or hematoxylin/eosin or oilred O/light green for lipid accumulation. Slides fixed inparaformaldehyde were rehydrated, incubated for 18 minutes in fresh oilred O, rinsed, and then incubated 1-2 minutes in fresh light green SFyellowish. The slides were then dehydrated, mounted, and the quantityand position of lipid deposition was analyzed by image analysis.

Quantitative Immunofluorescence (QIF)

Sections fixed in acetone were rehydrated in TBS+3% BSA for 30 minutes.The sections were incubated with primary antibody (anti-apo(a)immunosorbed on plasminogen, from Immunex, 1:1000 dilution; anti-totalTGF-beta BDA47, from R&D Systems, 1:200 dilution; MBPIIIB10₁anti-osteopontin antibody, from NIHDSHB, 1:200 dilution) in TBS+3% BSA.Sections were washed 3×3 minutes in PBS, then incubated withfluorescent-labeled second antibody for 2 hours. After washing 3×3minutes and mounting, bound fluorescence was quantitated by imageanalysis. Two markers could be examined on the same section usingfluorescein and rhodamine as distinct fluorescent labels with differentexcitation and emission characteristics.

Active TGF-beta was localized and quantitated following incubation ofslides with fluorescent-labeled extracellular matrix domain of theTGF-beta type II receptor (R2X), expressed in E. coli as aglutathione-S-transferase fusion protein.

Results

When confluent cells were injured in the presence of serum, many cellsmigrated into the wound area within 24 hours. Proliferation was alsostimulated under these conditions (7% of cells entered DNA synthesis,compared with 3% in an uninjured, control confluent culture). Theaddition of TGF-beta-1 (10 ng/ml) or tamoxifen (TMX; 10 μM) to rat cellsat the time of wounding substantially inhibited migration (approximately90% less cells crossed the boundary of the wound), consistent withprevious data that demonstrated that TGF-beta inhibited SMC migration inBoyden Chamber assays. The inhibition of migration by TMX was reversed(>90%) by a neutralizing antibody to TGF-beta-1 (25 μg/ml).

In contrast, TGF-beta and TMX did not significantly inhibit the entryinto DNA synthesis that was stimulated upon wounding. This observationis consistent with previous data that showed that TGF-beta and TMX slowSMC proliferation by extending the cell cycle in the G₂ phase, ratherthan by inhibiting or slowing entry into DNA synthesis.

These data agree with previous work that showed that apo(a) inhibitsTGF-beta activation in culture, thereby promoting SMC migration. Asdescribed in Example 4, apo(a) stimulated VSMC proliferation. Apo(a) isassociated with atherogenesis in man and in apo(a) transgenic mice. Whenapo(a) accumulates in conjunction with reduced levels of activeTGF-beta, both migration and proliferation will increase. TMX, whichstimulates formation of active TGF-beta, should ameliorateatherogenesis, regardless of whether migration or proliferation (orboth) play key roles in pathogenesis.

In adult rat aorta SMC, LDL accumulation is very low, both in freshlydispersed cell preparations and in primary and secondary cultures. Thisphenomenon is due to very low levels of LDL receptors (200-400receptors/cell), irrespective of whether the cells were exposed tolipoproteins.

In contrast, intimal SMC derived from rats 14 days after balloon injuryto the carotid artery have a greater (≈5 fold) uptake of LDL, due toincreased LDL receptor numbers (1500-2000 receptors/cell). When intimalcells or neonatal cells (displaying very similar properties) are treatedwith 10 ng/ml TGF-beta for 48 hours, these cells modulate, apparentlyirreversibly, to the adult phenotype. This phenotypic modulation isaccompanied by a down-regulation of LDL receptors (≈800 receptors/cell),with a reduction of LDL uptake of >80%. The presence of TGF-beta maytherefore reduce lipid accumulation by SMC.

The data obtained with apo(a) transgenic mice are consistent with thisprediction. In these mice, apo(a) is accumulated at high levels at theintimal surface of the aorta. TGF-beta activation is stronglydown-regulated from >80% in control aortas to <20% in apo(a) aortas.Lipid accumulation occurred at these sites in transgenic mice that werefed a lipid-rich diet and had elevated circulating LDL levels. Thus,reduced TGF-beta activity correlates with increased SMC accumulation ofLDL from the circulation. TMX, which is capable of elevating TGF-beta invivo, may inhibit lipid accumulation in vivo.

EXAMPLE 6 Effect of Idoxifene on Cultured Human VSMCs

Cultures of human VSMCs were prepared either by enzyme-dispersal usingcollagenase and elastase or using the explant technique in which cellsmigrate out from pieces of aorta (about 1 mm³) and proliferate,essentially as described in Example 3. Both enzyme-dispersed (ED) andexplant-derived (EX) cultures were prepared from the aortae of twoindividuals, and either EX or ED cultures were prepared from eightadditional donors. The two types of cultures have distinct morphologiesand growth characteristics. The EX cultures proliferated much morerapidly than the ED cultures. After six weeks of culturing both types ofcultures whenever confluence was attained, the total yield of cells wasapproximately 4 fold higher per gram wet weight of aorta in the EXcultures than the ED cultures. Consistent with this observation, the EDcultures had a longer population doubling time in DMEM+20% FCS (68±2hours; n=6) than the EX cultures (35±2 hours; n=6), p<0.001.

Idoxifene (IDX) is an analog of TMX which has been reported to haveenhanced anti-tumor activity (Chandler et al., Cancer Res., 51, 5851(1991); McCague et al., Organic Preparation & Proc. Int., 26, 343(1994)). The reduced side-effects of IDX compared with TMX and otherTMX-related analogs have prompted the selection of IDX for comparisonwith TMX. IDX at 5 μM inhibited increase in cell number by 30% and 28%(two EX cultures tested) compared to control, while cell growth in thepresence of 5 μM IDX and the neutralizing antibody to TGF-β (25 μg/ml)was 95+6% and 92±0% of control. In summary, both TMX and IDX inhibitedcell growth of EX-derived, but not ED-derived, hVSMCs to a similarextent (ED₅₀=5, 10 and 100 nM; n=3 experiments) and this effect wasreversible with the neutralizing antibody to TGF-β.

Despite the increasing use of animal models for vascular diseases, suchas transgenic mice and balloon-induced injury models, cell culturemodels of human VSMCs remain important tools because ofspecies-to-species variation. One problem associated with human cellculture models is the potential for variability in properties betweenindividuals due to gender and age, as well as genetic and environmentaldifferences. In this study, it was demonstrated that properties of VSMCcultures derived from ten different donors were very similar. The rateof proliferation, degree of differentiation indicated by expression ofthe contacile proteins SM-α-actin and SM-MHC and response to growthfactors of the smooth muscle cells were not influenced by the age or sexor genetic differences between the individuals.

By contrast, the method of establishing the VSMC culture had markedeffects on the properties of the cells. VSMCs derived by the explanttechnique had a spindle shaped morphology, proliferated rapidly(doubling time of about 35 hours) and lost expression of the contractileprotein SM-α-actin and SM-MHC in culture. VSMCs derived from the sameindividual by the enzyme-dispersal technique were larger, with stellatemorphology, proliferated more slowly (doubling time of about 68 hours)and retained high levels of expression of the contractile proteinsSM-α-actin and SM-MHC throughout many (>20) passages in culture. It istherefore important when comparing cell culture studies of human VSMCsto take into account the method used to establish the cultures.

The mechanisms which underlie the differences between the two types ofhuman VSMC culture were investigated. All of the differencesinvestigated the potential role of TGF-β result from production andactivation of TGF-β by the ED, but not EX cultures. Addition of aneutralizing antiserum to TGF-β to ED cultures altered the properties ofthe cells so that they resembled EX cells. Conversely, addition ofactive TGF-β to EX cells resulted in properties resembling ED cells.Furthermore, agents previously shown to inhibit rat VSMC proliferationby increasing TGF-β activity, such as TMX (Grainger et al., Biochem. J.,294, 109 (1993)) and heparin (Grainger et al., Cardiovas. Res., 27, 2238(1993)), inhibited the proliferation of EX but not ED cells.

A number of recent studies have demonstrated that reduced TGF-β activityis correlated with the development of atherosclerosis both in transgenicmouse models (Grainger et al., Nature, 370, 450 (1994)) and in man(Grainger et al., J. Cell. Biochem., 18A, 267 (1994)). The mechanismswhich control TGF-β production in the ED and EX human VSMC cultures maytherefore provide important clues as to the regulation of TGF-β activityin vivo. One possibility is that the VSMCs in the ED and EX culturescome from sub-populations of the VSMCs in the vessel wall which differin their ability to produce TGF-β. Evidence is accumulating forheterogeneity of VSMCs both in culture and in vivo and it will beinformative to determine whether equivalent sub-populations exist invivo by identifying a number of the genes which are differentiallyexpressed between the two types of culture.

If a reduction of TGF-β activity plays a role in atherogenesis, thenagents which elevate TGF-β activity, such as TMX, would be expected toreduce the incidence of myocardial infarction. The results describedabove indicate that TMX stimulates TGF-β production by human VSMC at10-100 fold lower concentrations than for rat VSMCs. Since TMX was shownto dramatically reduce the incidence of fatal myocardial infarction in arecent study of 1500 women (McDonald et al., Brit. Med. J., 303, 435(1994)), it is possible that an increase in active TGF-β, operating inan autocrine inhibitory loop, was responsible for these effects.

EXAMPLE 7 Tamoxifen Elevates TGF-β and Suppresses Diet-induced Formationof Lipid Lesions in Mouse Aortae

Treatment of Mice with TMX and Preparation of Aortic Sections

Adult (8-12 weeks old) male C57B16 mice in groups were weighed then fedad libitum either normal mouse chow (ICN/Flow), or a high fat dietcontaining 1.25% cholesterol, 7.5% saturated fat as cocoa butter, 7.5%casein and 0.5% sodium cholate, or high fat diet containing 15 μg TMXper gram, or high fat diet containing 1 μg TMX per gram. Water wasfreely available throughout. After three months on the respective diets,each mouse was re-weighed before sacrifice. The heart and attached aortawere embedded in Cryo-M-bed (Bright Instrument Co., Huntington, U.K.)and snap frozen in liquid nitrogen, then 4 μm frozen sections wereprepared as described previously (Paigen et al., Proc. Nat'l. Acad.Sci., 84, 3763 (1987); Paigen et al., Cancer Res., 45, 3850 (1985)).Platelet-poor plasma was prepared by adding blood taken at the time ofdeath to one tenth volume of 3.5% w/v trisodium citrate on ice. After 15minutes, the samples were spun (5,000×g; 15 minutes) and the plasmasupernatant retained. In the experiment with 4 groups of 15 mice, theplasma from 9 mice from each group was pooled for analysis of the lipidprofile of each group. Separate aliquots from the remaining 6 mice ineach group were stored at −20° C. until assayed.

Measurement of TGF-β in Plasma and Aortic Wall Sections

The (a+1)TGF-β in serum or platelet-poor plasma was measured by ELISA asdescribed above in Example 4. Active TGF-β was measured by ELISA usingtruncated extracellular domain of the type II TGF-β receptor (R2X).Active and (a+1)TGF-β were measured in 4 μm frozen aortic sections byquantitative immunofluorescence as described above in Example 4. ActiveTGF-β was measured using fluorescein-labelled R2X, (a+1)TGF-β wasmeasured using BDA19 antiserum (R & D Systems).

Analysis of Lipid Lesion Formation by Oil Red O Staining

For each mouse, 5 sections separated by 80 μm were fixed in 10% bufferedformalin, stained with oil red O and counter stained with light green asdescribed by Paigen et al., supra. The first and most proximal sectionto the heart was taken 80 μm distal to the point where the aorta becamerounded. The area of oil red O staining in each section was determinedwith a calibrated eyepiece, excluding lipid droplets less than 50 μm²,and the mean lesion area per section per mouse was calculated for eachmouse and each group of mice. Regions of focal lipid staining >500 μm²were defined as lipid lesions, and the number of such lesions persection per mouse was determined.

Lipoprotein Profile Analysis

One ml of pooled, platelet-poor plasma from each group of mice wasdiluted to 4 ml with buffer A (0.15 M NaCl, 0.01% (w/v) sodium EDTA and0.02% (w/v) sodium azide at pH 7.2) and ultracentrifuged at d=1.215 g/mlfor 48 hours at 4° C. 0.5 ml of the 2 ml lipoprotein fraction (d<1.215g/ml) was gel filtered through a sepharose 6B column by FPLC at roomtemperature. The column was eluted with buffer A at 0.4 ml/minute andfractions of 0.2 ml were collected and analyzed for cholesterol.Cholesterol was measured by the cholesterol oxidase method (SigmaDiagnostics) by adding 5 μl from each column fraction to 200 μl assayreagent in an ELISA plate (Maxisorp plates; Gibco). The assay plate wasincubated at 37° C. for 15 minutes and absorbance read at 492 nm. Serumfor calibration containing 200 mg/dL total cholesterol (SigmaDiagnostics) was used to convert absorbance readings to cholesterolconcentrations according to the manufacturer's instructions. Thepositions of elution of the major lipoprotein classes in mouseplatelet-poor plasma under the conditions described have been determinedpreviously (Yokode et al., Science, 250, 1273 (1990)). Fractions 1-9contained the very low density lipoprotein (VLDL), fractions 10 to 19contained LDL and fractions above 20 contained HDL.

Assays for Plasma Triglycerides, Cholesterol and Sex Hormones

Total plasma triglycerides was measured by the UV end-point glycerolkinase enzymatic method (Sigma Diagnostics). Total plasma cholesterolwas measured by the cholesterol oxidase method (Sigma Diagnostics)performed in ELISA plate wells as described above. 17-β-estradiol wasmeasured by a specific sandwich ELISA assay (Cascade Biochemicals) andtotal testosterone plus dihydrotestosterone by radio-immunoassay(Amersham International). All blood parameters (apart from thelipoprotein profile) were performed on six individual platelet-poorplasma aliquots in each group of mice.

Measurement of SM-α-actin and Osteopontin in Vessel Wall Sections

Four μm frozen sections were prepared from the heart/aorta blocksstained with oil red O for lipid lesions. One section adjacent to eachsection stained for lipid was stained for smooth muscle α-actin byquantitative immunofluorescence except that the mouse monoclonalantibody to smooth muscle α-actin, A-2547 (Sigma Chemical Co.), was usedas the primary antibody at 1:2000 dilution. Fluorescein-labelledanti-mouse IgG (Sigma Chemical Co.) was used as the second antibody at1:64 dilution. Osteopontin was measured in the next adjacent frozensection, using the mouse monoclonal antibody MBPIIIB 10 (NIHDevelopmental Studies Hybridoma Bank) labelled with biotin followed byfluorescein-labelled streptavidin.

Results

To determine the effects of TMX on TGF-β in the aortic wall and incirculation, an initial study was performed to establish an effectivedose. Adult (8 week old) male C57B 16 mice (a strain of mice susceptibleto lipid lesion formation on a high fat diet and which develop fattystreak lesions which resemble the early stages of atherosclerosis inman) in 3 groups were fed ad libitum for 28 days on either a normalmouse chow (low fat diet), or a high fat chow containing 0.5% sodiumcholate and 5% cholesterol (high fat diet), or high fat diet containing15 μg/g TMX (high TMX diet). The mice on the high TMX diet received anaverage of 1.1±0.3 mg/kg/day of TMX. Groups of 6 mice each were killedat intervals up to 28 days after starting the high TMX diet. ActiveTGF-β and active plus acid activatable latent TGF-β [(a+1)TGF-β] inserum samples and in the aortic wall were determined as described inExample 8. The (a+1)TGF-β increased detectably after 3 days reaching amaximum increase of 2.8-fold in serum and 10-fold in the aortic wall andcompared with control groups of mice on the high fat diet. After 7 days,(a+1)TGF-β in both the vessel wall and in serum declined slowly, so thatby 28 days, it was elevated by 2.4-fold in serum and 5.8-fold in theaortic wall. Active TGF-β also increased in response to the high TMXdiet and the kinetics of the initial increases in active TGF-β were verysimilar to those for (a+1)TGF-β, reaching a maximum at 7 days, with morethan 90% of the (a+1)TGF-β in serum and in the aortic wall was in theactive form at 7 days after starting the high TMX diet. However, between7 and 28 days, the increase in active TGF-β in both serum and in theaortic wall decline more rapidly than the (a+1)TGF-β so that after 28days, active TGF-β was only elevated by 1.5-fold in serum and 2.2-foldin the aortic wall. The decrease in the proportion of active TGF-β after7 days appears to be due to the induction of plasminogen activatorinhibitor-1.

In a further experiment, adult (8 week old) C57B16 mice in 3 groups of15 were fed on the diets described above, together with a fourth groupof 15 mice fed a high fat diet containing 1 μg/g TMX (low TMX diet). Themice on the high TMX diet received an average dose of 1.1±0.3 mg/kg/dayof TMX on the low TMX diet received 0.08±0.02 mg/kg/day. The remainingmice were killed after 3 months on the diets and the heart, lungs andaortae were embedded and snap-frozen in liquid nitrogen. Platelet-poorplasma was prepared from a terminal bleed. None of the mice in the 4groups showed anatomical abnormalities, although the mice fed TMX at thehigh or low doses gained less weight during the period of the experimentthan the mice on either the low fat or high fat diet (Table 2). Theconcentrations of both active and (a+1)TGF-β in plasma and in the aorticwall were significantly increased by the high TMX diet. On the low TMXdiet, only the active TGF-β in plasma did not show a significantincrease (Table 2). The effects of TMX on TGF-β after 3 months of thehigh TMX diet were significantly lower than in mice treated for 28 days.

TABLE 2 Effects of High Fat Diet and Tamoxifen on C57B16 Mice Low FatHigh Fat Low TMX High TMX TMX — — 0.08 ± 0.02 1.1 ± 0.3  (mg/kg/ day)Weight 8 ± 2 9 ± 1  5 ± 2**  2 ± 1*** gain over 3 months (g) (a + 1 )TGF-β Plasma 11 ± 4  12 ± 3   18 ± 5** 22 ± 6*** (ng/ml) Vessel 22 ± 4 20 ± 2   32 ± 4** 44 ± 8*** Wall (arbitrary units) Active TGF-β Plasma 8± 3 8 ± 2 10 ± 3  12 ± 3*** (ng/ml) Vessel 20 ± 13 18 ± 4   28 ± 3** 33± 5*** Wall (arbitrary units) Lesions 0.7 ± 0.1  3.6 ± 1.0*  2.6 ± 0.8** 1.1 ± 0.3*** per mouse^(a) Lesion 230 ± 50  6860 ± 4660 ± 823 ± area/1480* 960** 220*** section/ mouse (μm²) 17β- 0.28 ± 0.10 0.39 ± 0.140.40 ± 0.20 0.25 ± 0.08  estradiol (ng/ml) Total 16 ± 2  14 ± 3  13 ± 5 11 ± 7   Testoster- one (ng/ml) Total 71 ± 2  92 ± 4*  79 ± 3** 83 ±4*** Plasma Choles- terol (mg/dl) VLDL 4 30 38 42 Choles- terol (mg/dl)LDL 8 33 27 27 cholesterol (mg/dl) HDL- 58  27 11 14 cholesterol (mg/dl)Total 142 ± 15  109 ± 5*  111 ± 9  204 ± 36*** Tri- glycerides (mg/dl)SM-α- 146 ± 6  138 ± 8  168 ± 14  204 ± 12*** actin (arbitrary units)Osteo- 2 ± 1  46 ± 16* 30 ± 11  5 ± 3*** pontin (arbitrary units)

Serial sections from the aortic sinus region were analyzed for lipidlesions using the oil red O staining protocol and sectioning strategy asdescribed by Paigen et al., supra. Small regions of luminal lipidstaining were detected in mice on the low fat diet, but most of thevessel wall was devoid of lipid deposits in this group. In mice fed thehigh fat diet, there was a 5-fold increase in the number of lipidlesions in the aortic wall but in the mice fed the TMX diets, there wasa dose-dependent decrease in the number of lesions with a 86% decreaseof diet-induced lesions on the high TMX diet (Table 2). The aortic wallarea stained with oil red O was measured for each group of mice. Mice onthe high fat diet had lesion areas (per section per mouse) of 6860±1480μm² (n=15) consistent with previous published results (Emerson et al.,Am. J. Path., 142, 1906 (1993); Paigen et al., Arteriosclerosis, 10, 316(1990)). The high TMX diet and low TMX diets reduced the lesion areas by88% (n=15; p<0.001) and 32% (n=15; p<0.01) respectively (Table 2). TMXtherefore causes a dose-dependent inhibition of diet-induced lipidlesions in C57B16 mice.

High or low TMX diets significantly lowered total plasma cholesterol byapproximately 10% compared with mice on the high fat diet. Analysis ofthe lipoprotein profiles showed that for the mice on the low fat diet,most of the cholesterol was in the HDL fraction. After 3 months on thehigh fat diet, however, there was a marked increase in very low densitylipoprotein (VLDL) cholesterol of approximately 7-fold (Table 2) and LDLcholesterol (4-fold) whereas the amount of cholesterol in the HDLfraction was reduced by approximately 50% (Table 2). The high and lowTMX diets had only small effects on the amount of cholesterol in VLDL orLDL, but further reduced the HDL cholesterol by approximately 50% (Table2), accounting for most of the overall reduction in cholesterol. Incontrast to the decrease in total plasma cholesterol concentrationcaused by the high TMX diet, there was an increase in plasmaconcentration of triglyceride (Table 2).

The high or low TMX diets did not affect the very low plasmaconcentrations of 17β-estradiol in the male mice (Table 2). The meantotal testosterone concentration (assayed as testosterone plusdihydrotestosterone) was not significantly altered by the TMX diets,although the range of testosterone concentrations was larger than in themice on the high fat diet, suggesting that TMX may affect testosteronelevels in individual mice. However, it is unlikely that changes in thelevels of the primary sex hormones in response to TMX are responsiblefor the inhibition of lipid lesion formation. Medial smooth muscle cellsin transgenic apo(a) mice which expressed osteopontin, a marker ofde-differentiated smooth muscle cells, are the site of focal apo(a)accumulation and very low TGF-β activity. The accumulation ofosteopontin occurred in mice on a low fat or high fat diets and wastherefore independent of the accumulation of lipid at the sites of lowTGF-β activity. In the C57B16 mice fed the high fat diet, sectionsadjacent to the lipid lesions identified by oil red O staining showedregions of high osteopontin accumulation, whereas there was almost noosteopontin accumulation in the aortic sections from mice on the highTMX diet. The type(s) of cells in the aortic wall (e.g., VSMCs,macrophages, etc.) from which the osteopontin was derived, were notidentified. Similar experiments in which the accumulation of smoothmuscle α-actin was assayed showed an inverse pattern to that forosteopontin. There were regions of low SM-α actin expression in adjacentsections to lipid lesions, whereas the amount of SM-α actin wasincreased in the sections from mice on the high TMX diet. Similarresults to those described above for C57B16 mice have been observed inthe transgenic apo(a) mouse when these mice were fed a high fat diet.That is, both the lesion areas and number of lesions for both strains ofmice were reduced by approximately 90%.

This example demonstrates that TMX strongly inhibits the formation oflipid lesions induced by a high fat diet in a susceptible strain ofmice. The data show that a major effect of TMX in the C57B16 mice is toelevate TGF-β in aortic wall and in circulation. This is consistent withprevious evidence that TMX increases the production of TGF-β by VSMCsand other types of cells in vitro and in breast tumor cells in vivo. Thesuppression of osteopontin accumulation and the increase in SM-α actinin mice treated with TMX is consistent with previous observations on theapo(a) transgenic mouse (Example 4). These mice showed largeaccumulations of osteopontin at sites where focal accumulations of highconcentrations of apo(a) result in decreased TGF-β activity in thevessel wall. The activation of the smooth muscle cell was also marked bya decrease in local SM-α actin concentration and occurred in the mice ona low fat diet in the absence of lipid accumulation. On a high fat diet,lipid accumulation occurred at the sites of apo(a) accumulation andlesions formed in two stages: activation of the VSMCs as a result of lowTGF-β activity and subsequently uptake of lipid by the activated cellswhen the mice are subjected to a high fat diet. Thus, the cardiovascularprotective effect of TMX in mice may be due to elevation of TGF-β in theartery wall which prevents VSMC activation and consequently inhibitslipid accumulation on a high fat diet. TMX causes an overall 2-foldincrease in active TGF-β in the aortic wall in C57B16 mice and a similarincrease in apo(a) transgenic mice would restore the overall TGF-βconcentration to that observed in normal littermate mice lacking theapo(a) gene. This hypothesis therefore predicts that TMX would preventlipid lesion formation in apo(a) mice on a high fat diet. It is ofinterest that the cardiovascular protective effects of TMX againstdiet-induced lipid lesions in mice reported here were obtained at dosessimilar to those used in breast cancer therapy.

EXAMPLE 8 Determination of Active and Acid Activatable TGF-β in HumanSera, Platelets and Plasma by Enzyme-Linked Immunosorbent Assays

Antibodies

The antibodies to TGF-β used for the ELISAs were BDA19 (a chickenpolyclonal IgY antibody which neutralizes TGF-β activity) and BDA47 (anaffinity purified rabbit polyclonal IgG antibody), both obtained fromR&D Systems (Oxford, U.K.). Goat anti-rabbit IgG coupled to horseradishperoxidase was obtained from Sigma Chemical Co. (Poole, U.K.). TGF-βstandards were obtained from Peninsula (St. Helens, U.K.; purifiedporcine TGF-β1) and Amersham International (Amersham, U.K.; recombinanthuman TGF-β1). To refer the ELISA data obtained with these TGF-β1s tothe interim international standard, bovine TGF-β1 (89/516) was obtainedfrom the National Institute of Biological Standards and Control (PottersBar, U.K.). TGF-β2 and TGF-β3 isoforms were obtained from R&D Systems).The TGF-β standards were dissolved in 25 mM Tris/HCl pH 7.4 containing50 μg/ml fatty acid free bovine serum albumin (FAF-BSA) to give 5 μg/mlstock solutions. The concentration of the standard TGF-β solutions waschecked against the bioassay of DNA synthesis in MvLu epithelial cells(see below). Both TGF-β standards gave an ED₅₀ for inhibition of DNAsynthesis in the MvLu bioassay of between 2-3 pM which agrees well withthe previously reported value of 2 pmol/L (Danielpur et al., J. CellPhysiol., 138, 79 (1989)).

Growth Factors

Platelet-derived growth factor (PDGF) AA and BB homodimers and epidermalgrowth factor (Bachem Inc., Saffron Walden, U.K.) were dissolved in 25mmol/L Tris/HCl, pH 7.4 containing 1% FAF-BSA to give 0.3 μmol/L stocksolutions. Basic fibroblast growth factor (0.56 μmol/L) interleukin 1 β(0.59 μmol/L), transforming growth factor α (1.81 μmol/L), interferon γ(0.59 μmol/L) and insulin-like growth factor I (0.59 μmol/L; all fromBachem Inc.) were dissolved in sterile MilliQ water to give stocksolutions of the concentrations indicated. Angiotensin II and endothelinI (Sigma Chemical Co.) were dissolved in sterile MilliQ water to give 10μmol/L stock solutions.

Recombinant Expression of the TGF-β Type II Receptor

The extracellular domain of the TGF-β type II receptor was amplifiedfrom the vector H2 3FF (Lin et al., Cell, 68, 775 (1992)) using thepolymerase chain reaction (PCR). The vector DNA was linearized with NotI, precipitated and resuspended at 10 ng/μL. Amplification was carriedout in a 50 μl reaction containing 2.5 μl DNA, 5 μl 10×TAQ buffer (LKBPharmacia; Upsalla, Sweden), 250 ng of each oligonucleotide primer(GAATTCCCATGGGTCGGGGGCTGCTC (SEQ ID NO:1) and GAATTCGTCAGGATTGCTGGTGTT(SEQ ID NO:2); Wellcome Protein and Nucleic Acid Chemistry Facility,University of Cambridge), 1 U TAQ polymerase and a mixture of dATP,dTTP, dCTP and dGTP to give a final concentration of 200 μM for eachnucleotide. The sample was overlaid with 50 μL paraffin oil. Thereaction was carried out using a thermal cycler (PREM; Cambridge, U.K.)for 30 cycles (denaturing at 94° C. for 1 minute, annealing at 55° C.for 2 minutes, elongation at 72° C. for 2 minutes). The 450 bp fragmentproduced was purified by electrophoresis in low gel temperature agarose,digested with EcoRI and cloned into the glutathione-S-transferase fusionvector pGEX 2T (LKB Pharmacia). Vectors carrying inserts in the requiredorientation were identified by plasmid mapping. The sequence of theinsert was checked by subcloning the 450 bp EcoRI fragment from thechosen clone (pGT1C) into Bluescript KS+ followed by double strandsequencing. The sequence showed a single base change (C to A atposition+13 from the initiation codon) compared to the publishedsequence (Lin et al., supra.) which introduces a leu to met mutation inthe protein.

Protein Purification

An overnight culture of E. coli TG1 containing pGT1C was diluted 1:100into fresh 2YT medium (500 mL) containing 270 μmol/L ampicillin andgrown to an OD₆₀₀ of 0.5. Production of the fusion protein was inducedby addition of 1 mM isopropylthiogalactoside and the cells wereharvested 5 hours later by centrifugation. The bacteria were resuspendedin 50 mL phosphate buffered saline (PBS; 150 mmol/L NaCl, 2 mmol/LNa₂HPO₄, 4 mmol/L Na₂HPO₄, pH 7.3) containing 1% Triton X-100 and 1mmol/L PMSF and lysed by sonication for 5 minutes. The lysate wascentrifuged (10,000×g; 5 minutes) and the fusion protein was purifiedfrom the supernatant by the one step purification method of Smith andJohnson (Gene, 67, 31 (1988)). FPLC of the purified glutathione-bindingproteins on a Superdex 200 HR column in 20 mM ammonium bicarbonate, pH8.0, demonstrated that >95% of the protein present was the desired 43kDa TGF-β receptor fusion protein.

ELISA to Measure Total TGF-β

Maxisorp 96 well ELISA plates (Gibco; Uxbridge, U.K.) were coated withthe capture antibody by incubating with 50 μL BDA19 anti-TGF-β chickenIgY (40 μg/mL) diluted in Tris-buffered saline (TBS; 137 mmol/L NaCl, 50mmol/L Tris/HCl, pH 7.4) and shaking the plates until dry by evaporationat room temperature (approximately 12 hours). The plates were washed 3×3minutes with PBS, blocked with 350 μL 3% FAF-BSA in TBS for 1 hour,washed 3×3 minutes with TBS and incubated for 2 hours with 100 μL oftest samples or dilutions of a TGF-β stock solution for calibration. Thepurified porcine TGF-β stock solution. diluted in TBS to concentrationsbetween 0.4 pmol/L and 4000 pmol/L was used for calibration unlessotherwise indicated.

The plates were washed (3×3 minutes) with TBS+3% FAF-BSA+0.1% TritonX-100 (wash buffer) and incubated with 20 μL detection antibody (BDA47;anti-TGF-β (rabbit IgG)) at 1 μg/mL in wash buffer for 1 hour. Theplates were rinsed with wash buffer (3×3 minutes) and incubated with anantibody against rabbit IgG conjugated to horseradish peroxidase (SigmaA-6154) at 1:2500 dilution in wash buffer for 1 hour. After washing (3×3minutes with wash buffer), the plates were incubated for 15 minutes withthe chromogenic substrate orthophenylenediamine (Sigma) according to themanufacturer's instructions. The reaction was stopped by addition of anequal volume of 3M HCl and the absorbances read on an ELISA plate reader(Titertek Multiscan; Flow Laboratories, High Wycombe, U.K.) within 15minutes of stopping the reaction. Absorbances were converted intoquantities of TGF-β protein using the calibration curve from the TGF-βstandard.

ELISA to Measure Active TGF-β

This ELISA was performed as for the ELISA to assay total TGF-β except:(i) the ELISA plates were coated with the purified TGF-β receptor fusionprotein using 20 μL of a 50 μg protein per mL of solution in TBS and(ii) the detection reagent (BDA47) was used at 5 μg/mL.

Mink Lung Epithelial DNA Synthesis Bioassay

Mink lung epithelial cells (MvLu; American Type Culture Collection;passage 49-60) were subcultured at 1:5 dilution in DMEM+10% FCS. After24 hours, the medium was replaced with DMEM+10% FCS containing thesample (<1% v/v) or standards in the presence and absence ofneutralizing antiserum to TGF-β (BDA19) at 10 μg/ml. DNA synthesisduring a 1 hour pulse of 6-[³H]-thymidine (5 μCi/ml; AmershamInternational) was determined 23 hours after addition of test medium.TGF-β activity was calculated as the proportion of the inhibition of DNAsynthesis which was reversed in the presence of neutralizing antibody,using a standard curve to convert the inhibition values into quantitiesof active TGF-β. Purified porcine TGF-β diluted in TBS was used as thestandard unless otherwise indicated.

Preparation of Conditioned Culture Media, Human Platelets, Platelet-PoorPlasma and Serum

Medium (DMEM+20% FCS) was conditioned for 24 hours on cultures of adulthuman aortic VSMCs obtained by enzymatic dispersion of aortic media asdescribed above.

Twenty mL of peripheral venous blood was collected from 12 healthy malevolunteers (aged 23-54); 10 mL were aliquoted immediately into tubescontaining 1.1 mL of sterile 3.8% (w/v) trisodium citrate in MilliQwater at room temperature. The samples were centrifuged (250×g; 15minutes) to remove red blood cells. Apyrase (Sigma) was added to theplatelet-rich plasma to a final concentration of 100 mg/L to preventplatelet degranulation; PMSF (1 mmol/L) and aprotinin (1 mg/L) wereadded to prevent proteolytic activation or degradation of TGF-β. Thesesamples were centrifuged (700×g; 15 minutes) and the supernatantplatelet-poor plasma was separated from the platelet pellet. Theplatelet-poor plasma was kept at room temperature until assayed byELISAs within 2 hours of preparation or was stored in 0.5 mL aliquots at−80° C. The platelet pellet was resuspended in 10 mL (i.e., the originalvolume of blood) of a buffered saline solution (145 mmol/L NaCl, 5mmol/L KCl, 10 mmol/L glucose, 10 mmol/L MgSO₄, 0.5 mmol/L EGTA, 1mmol/L PMSF, 1 mg/L aprotinin, 10 mmol/L HEPES, pH 7.4) andrecentrifuged as before. The washed platelet pellet was resuspended in10 mL of buffered saline solution and the platelet concentration wasdetermined by hemocytometer. Platelets were lysed by ultrasonicationuntil <10% of unlysed platelets were detected by hemocytometer. Humanplatelet suspensions were also obtained form the Blood TransfusionService, Cambridge, U.K. The platelets were collected by centrifugation(3,000×g; 3 minutes) and approximately 0.1 g of platelets wereresuspended in 0.5 mL MilliQ water and lysed by three cycles offreeze-thawing. The membrane fragments were removed by centrifugation(14,000×g; 10 minutes) and the supernatant was mixed with an equalvolume of 2×TBS.

The remaining 10 mL of freshly drawn blood samples were dispensedimmediately into polypropylene tubes and allowed to clot at roomtemperature for 2 hours. The clotted samples were centrifuged (1,000×g;4 minutes), the serum was removed and either stored on ice until assayedwithin 2 hours or stored at −80° C. until assayed. The clot was washedthree times by centrifugation (1000×g; 4 minutes) in 5 mL of 150 mMphosphate buffer, pH 7.0, and the third wash was retained for TGF-βassays. The washed clot was dissolved in 5 mL of 150 mM phosphatebuffer, pH 2.0, for 30 minutes, then neutralized by addition of 5 mL of150 mM phosphate buffer, pH 12.0. The samples were assayed for TGF-βimmediately or stored in 1 mL aliquots at −80° C.

All blood-derived samples, stored at −80° C., were not thawed untilassayed. The initial freeze-thaw cycle resulted in less than 10% loss oftotal or active TGF-β activity in the ELISAs. However, three additionalfreeze-thaw cycles of samples containing TGF-β in active or latent formwas sufficient to cause loss of approximately 90% activity.

Bioassays of PDGF

PDGF was bioassayed by its mitogenic activity on human VSMCs derived byexplant as described previously (Kocan et al., Methods in Cell Biology,eds. Harris, C. C., Trump, B. F., and Stenes, G. D., Academic Press(1980)). VSMCs were made quiescent by incubation in serum-free DMEM for48 hours. Samples of serum or platelet-poor plasma were added at a finalconcentration in DMEM of 5% or 20%, respectively. DNA synthesis wasassayed by [³H]-thymidine (Amersham International; 5 μCi/mL)incorporation between 12 hours and 36 hours after addition of the testsamples to the cells. The proportion of DNA synthesis due to PDGF wasestimated by the addition of polyclonal antibody (50 mg/L) whichneutralizes all forms of PDGF to replicate cell samples.

Results

An ELISA was set up to detect total (α+1) TGF-β using the polyclonalchicken IgY antibody BDA19 as the capture reagent. The assay detectedpurified porcine TGF-β in TBS in the range of 4 pmol/L to 2000 pmol/Lwith half-maximal change in absorbance (ΔA_(50%)) of 280±80 pmol/L(n=7). Using recombinant human TGF-β1 in TBS, the assay detected TGF-βin the range 8 pmol/L to 2000 pmol/L with a ΔA_(50%) of 320±120 pmol/L(n=3). Direct comparison of the TGF-β1 (R&D Systems) was made with theinterim international bovine TGF-β (89/516). An ampoule of 89/516containing 1500 units (approximately 80 ng protein; 32 pmol) wasdissolved in sterile water to 800 μl and serially diluted in TBS andsimilar dilutions of the R&D Systems TGF-β1 made. Comparison of thecalibration curves showed that a nominal 1.0 pmol at R&D TGF-β1 had anactivity of 130±8 units. To test the specificity of the capture antibodyin the total TGF-β assay, it was replaced with nonimmune chicken IgY(R&D Systems). The change in absorbance in the presence of 4000 pmol/Lof purified porcine TGF-β1 was less than 5%, indicating that TGF-βbinding under the assay conditions was specific to the capture agent.

To test whether the ELISA detected acid activatable, latent forms ofTGF-β, a sample of human platelets from the blood bank was lysed andassayed before and after activation of the TGF-β (Wakefield et al., J.Biol. Chem., 263, 7646 (1985); Assoian et al., J. Cell Biol., 102, 1031(1986)). The latent TGF-β was converted to active TGF-β by addition of5% vol/vol 150 mmol/L sodium phosphate buffer at pH 2.0 for 5 minutes,then neutralized by addition of 5% vol/vol 150 mmol/L sodium phosphatebuffer at pH 12.0 (Barnard et al., Biochim. Biophys. Acta, 1032, 79(1990)). Control samples were treated with 10% vol/vol 150 mmol/L sodiumphosphate buffer at pH 7.0. The MvLu cell bioassay of the untreated andacid-treated platelet lysate showed that the amount of active TGF-β wasincreased 5.1-fold after acid activation of the latent TGF-β, indicatingthat approximately 80% of the TGF-β present in the unactivated samplewas in the acid activatable, latent form. When assayed by the totalTGF-β ELISA, the control aliquot contained 680±80 pmol/L TGF-β (n=3) byELISA and the acid-activated aliquot contained 600±120 pmol/L TGF-β(n=3). These results show that the total TGF-β ELISA does notdistinguish between active and acid activatable TGF-β from humanplatelets.

The precise conditions for activation of the small and large complexesof latent TGF-β have not been characterized and there is some evidencefor the existence of two pools of latent TGF-β which differ in theconditions required for activation. Therefore, TGF-β is defined as thatpool of latent TGF-β which is acid-activatable by the treatmentdescribed above (i.e., exposure to pH 2.0 for 5 minutes beforeneutralization to pH 7.0 without overshoot). Longer exposure to pH 2.0did not significantly affect the concentration of activated TGF-β and itremains to be determined which form(s) of latent TGF-β are activatedunder the defined conditions.

A second ELISA was established to measure active TGF-β in the presenceof latent TGF-β using a truncated TGF-β type II receptor protein fusedto glutathione-S-transferase as the capture reagent. This assay detectedpurified porcine TGF-β1 in TBS in the range of 20 pmol/L to 4000 pmol/Lwith a ΔA_(50%) of 680±160 pmol/L (n=4) and recombinant human TGF-β1 inTBS in the range of 40 pmol/L to 4000 pmol/L with a ΔA_(50%) of 720±120pmol/L (n=3). To test the specificity of the truncated receptor fusionprotein as the capture agent, it was replaced withglutathione-S-transferase. The change in absorbance in the present of4000 pmol/L of purified porcine TGF-β1 was less than 5%, indicating thatTGF-β binding was specific to the capture agent under the assayconditions.

To confirm that the active TGF-β ELISA did not detect acid activatable,latent TGF-β, samples of human platelet TGF-β before and after acidactivation were assayed. The active TGF-β ELISA gave 160±40 pmol/L (n=3)in the unactivated sample and 640±80 pmol/L (n=3) TGF-β in theacid-activated sample, consistent with the data obtained from the (a+1)TGF-β ELISA and the MvLu cell bioassay described above. The ability ofthe ELISA to discriminate between active and latent TGF-β was furtherdefined in studies on TGF-β in fresh human platelets (see below).

To test the reproducibility of both ELISAs, 24 aliquots of a sample oflysed human platelets from the blood bank was assayed simultaneously byboth assays. The value for active TGF-β was 200 pmol/L with acoefficient of variation of 7.4% and the corresponding value for (α+1)TGF-β was 640 pmol/L with a coefficient of variation of 6.8%. Furtheraliquots of the same platelet lysate were also analyzed blind by fourindependent operators using both ELISAs on eight separate occasions. Theinter-assay coefficient of variation was 13.2% for the active TGF-βassay and 12.2% for the (α+1 ) TGF-β assay.

The relative sensitivity of each ELISA to the three isoforms of TGF-βwas determined. Recombinant human TGF-β1, TGF-β2 and TGF-β3 (400 pmol/L)in TBS were assayed using each ELISA, expressing the absorbance forTGF-β2 and TGF-β3 as a percentage of the absorbance for TGF-β1. BothELISAs detect TGF-β1 and TGF-β3 with similar sensitivity, but TGF-β2 wasdetected with approximately 10-fold less sensitivity than the otherisoforms in the (α+1) TGF-β ELISA and 100-fold less sensitivity in theactive TGF-β ELISA. The relative sensitivities for the isoforms in theactive TGF-β ELISA are qualitatively consistent with the relative TGF-βisoform affinities of the type II TGF-β receptor (Massagué, Ann. Rev.Cell Biol., 6, 597 (1990)). The slightly greater relative sensitivity ofthe active TGFβ ELISA to TGF-β3 than the (α+1) TGF-β ELISA would resultin an overestimate of the proportion of active TGF-β in a sample whichwas composed mostly of TGF-β3 if the assays were calibrated using aTGF-β1 standard. The proportion of active TGF-β in samples containingonly the TGF-β2 isoform cannot be determined accurately by these ELISAsat concentrations below 4000 pmol/L. The concentration of TGF-β2 inhuman serum has been reported as <5 pmol/L (Danielpur et al., AnnalsN.Y. Acad. Sci., 593, 300 (1990)).

The cross-reactivity of both ELISAs to a variety of other peptide growthfactors was determined at concentrations which have a maximal biologicaleffect in cell culture. Neither assay gave a change of greater than 5%in absorbance in response to PDGF-AA (3.3 nmol/L), PDGF-BB (3.3 nmol/L),basic fibroblast growth factor (5.6 nmol/L), epidermal growth factor(15.9 nmol/L), insulin-like growth factor I (1.3 nmol/L), angiotensin II(100 nmol/L), endothelin I (100 nmol/L), interleukin 1β (588 pmol/L),transforming growth factor α (1.8 nmol/L), or interferon γ (588 pmol/L).

There are several reports that TGF-β binds to serum components andextracellular matrix components with high affinity. For example,McCaffrey and co-workers demonstrated that TGF-β associatesnon-covalently with the major serum protein, α2-macroglobulin (J. CellBiol., 109, 441 (1986)). However, preparation of the TGF-β standardsolutions in the presence of 1.4 μmol/L human α2-macroglobulin or 10%FCS did not affect the ΔA_(50%) by more than 10% compared with theΔA_(50%) for the standard TGF-β solutions diluted in TBS in eitherELISA. Therefore, any non-covalent interactions formed between TGF-β andα2-macroglobulin or with components of FCS do not prevent active TGF-βfrom binding to the type II TGF-β receptor in the active TGF-β ELISA orto the capture antibody in the (α+1) TGF-β ELISA, nor do they inhibitbinding by the detection antibody. It has been noted in a previousreport that purified TGF-β and α2-macroglobulin may not interact in thesame way as endogenous serum TGF-β and α2-macroglobulin(O'Conner-McCorua et al., J. Biol. Chem., 262, 14090 (1987)).

The active TGF-β concentration was measured in three samples of medium(DMEM containing 10% FCS) conditioned for 24 hours on human VSMCs whichproduce active TGF-β. The values obtained with the active TGF-β ELISAwere compared with those obtained using the MvLu cell bioassay (Table3).

TABLE 3 Active TGF-β concentration in medium conditioned on human VSMCsActive TGF-β (pM) Sample MvLu Assay Active TGF-β ELISA 1 584 ± 24 552 ±32 2 356 ± 32 400 ± 24 3 488 ± 40 484 ± 16 The amount of active TGF-βpresent in three different samples of DMEM + 20% FCS which had beenconditioned on human VSMC cultures for 24 hours was determined inquadruplicate using the DNA synthesis bioassay in MvLu epithelial cellsand the active TGF-β ELISA.

The results obtained by the two assays were not statistically differentfor any of the three samples tested (p=0.88, 0.48 and 0.99, usingstudents unpaired t-test). Thus, the ELISA gives values for active TGF-βconcentrations in conditioned medium which are closely consistent withthe MvLu cell bioassay used previously. Where possible, it is importantto demonstrate consistency between the active TGF-β ELISA and thebioassay for conditioned media and other biological fluids. For example,it has recently been reported that direct addition of conditioned mediato ELISA microwells can lead to inaccurate measurement of TGF-β forreasons that are not fully understood (Danielpur, J. Immunol. Methods,158, 17 (1993)). Protocols which activate and concentrate TGF-βs topartially purify the samples and exchange the buffer were recommended(Danielpur, supra).

Another factor which might interfere with the assays is any peroxidasespresent in serum which bind to the capture reagents. To test forperoxidases, the capture antibody in the (α+1) TGF-β assay was replacedwith non-immune chicken IgY, and the truncated receptor fusion proteinin the active TGF-β assay was replaced with glutathione-S-transferase.The change in absorbance in either assay was less than 5% in thepresence of either DMEM containing 10% FCS or human serum from donors A,E, K, or N in Table 5. These data indicated that any peroxidase activityin FCS or human serum did not significantly affect the assays of (a+1)or active TGF-βs.

TABLE 4 Active and (α + 1) TGF-β concentrations in human sera TGF-β(pmol/L) Unactivated serum Acid-activated serum Donor Active (α + 1)Active (α + 1) A <40 240 240 240 B 120 120 120 120 C 200 320 320 320 D240 240 240 240 Serum samples from four male donors were assayed in asingle experiment for active and total TGF-β by the ELISAs before andafter acid activation. All samples were assayed in quadruplicate.

The above experiments suggested that the ELISAs could be used to measureTGF-β in human serum and the use of the assays for sera was thereforecharacterized. It was found that the calibration curves for both theactive and (α+1) TGF-β assays were not affected when purified porcineTGF-β was added to human serum (donor E in Table 5) which contained verylittle TGF-β by either ELISA.

TABLE 5 (α + 1) and active TGF-β concentrations in human serum SamplesTGF-β (pmol/L) Donor Active (α + 1) % active E <20 <4 — F <20 <4 — A <20240 <8 G 20 80 25 H 80 80 100 I 80 80 100 J 80 120 66 K 160 1120 14 C280 320 88 L 320 320 100 M 360 320 113 N 1400 1400 100 Serum samplesfrom 12 male donors aged between 23 and 54 were assayed immediatelyafter preparation for active and (α + 1) TGF-β by the ELISAs described.All samples were assayed in quadruplicate by each ELISA in a singleexperiment.

For human sera comparisons of active TGF-β concentrations by the ELISAand the MvLu cell bioassay were not possible because human seruminhibited MvLu DNA synthesis by a mechanism independent of TGF-β. Thepresence of 10% (v/v) serum from any of 4 donors (A, H, J, and K inTable 5) inhibited DNA synthesis in MvLu cell cultures by more than 95%.This inhibition was not reversed by the presence of neutralizingantibodies to TGF-β, indicating that the human sera contained aninhibitor of DNA synthesis in MvLu cells which masked any effect ofTGF-β. The MvLu cell bioassay cannot therefore be used to determine theconcentration of active TGF-β in unfractionated human serum samples.

Alternative approaches were therefore required to validate the ELISAassays for direct use with human serum. The main requirement was todetermine whether human sera contain non-TGF-β components whichsignificantly affected the TGF-β concentrations estimated by eitherassay. Overestimated values of TGF-β would be obtained if a serumcomponent was bound specifically or nonspecifically by the capture agentin either assay and was also recognized by the detection antibody or bythe antibody to rabbit IgG linked to horseradish peroxidase.Alternatively, underestimated values would result if a serum componentcompeted with TGF-β for the capture agent in either assay but was notrecognized by the detection antibody. In a previous study in which TGF-βin unfractionated serum (after transient acidification) was determinedby a radio-receptor assay, it was found that components in the seruminterfered with the assay (O'Connor-McCourt et al., J. Biol. Chem., 262,14090 (1987)). This resulted in a dilution curve which was not parallelto the standard dilution curve and estimates of TGF-β were 20 to 40times lower than those obtained by acid-ethanol extraction of the samesamples. Thus, it is possible that serum components which result ineither overestimated or underestimated TGF-β values in our ELISAs wouldalso interfere with other assays (receptor binding orradio-immunoassays) used to validate serum TGF-β concentrationsestimated by the ELISAs. Therefore, a more rigorous test for interferingcomponents in serum was required. This was achieved by determiningwhether the concentrations of active and (α+1) TGF-β concentrations insera were internally consistent before and after activation of latentTGF-β by acid treatment. Only under very implausible circumstances wouldconsistent accounting of active and (α+1) TGF-β be obtained in thepresence of serum components which interfered with either or bothassays.

ELISAs of (α+1) and active TGF-β concentrations were performed on thesera from 4 male donors before and after the sera were acidified to pH2.0 and neutralized to pH 7.0 as described for the lysed human plateletsamples. For each of the sera in Table 4, there was no difference withinthe accuracy of the assays between the amount of (α+1) TGF-β before andafter acid treatment. Furthermore, after acid treatment, the amount ofactive TGF-β was not significantly different from the amount of (α+1)TGF-β. These results imply that it is very unlikely that the sera testedcontained components which interfered with either TGF-β ELISA since theywould cause significant imbalances in the quantitative accounting of theamounts of active and (α+1) TGF-β before and after acid treatment. Theuse of acid treatment of the sera and reassay of the active and (α+1)TGF-β concentrations therefore provides an important internal controlfor the TGF-β assays when used directly for sera or complex biologicalfluids.

The sera from 12 male donors (aged 23 to 54) were assayed for active and(α+1) TGF-β by the ELISAs (Table 5). The mean (α+1) TGF-β concentrationwas 330 pmol/L, but the variation was very large (range less than 4pmol/L to 1400 pmol/L). Similarly, the mean active TGF-β concentrationwas 230 pmol/L, and the range was from less than 20 pmol/L to 1400pmol/L. The proportion of the (α+1) TGF-β present which was activeranged from <10% to 100% with a mean of 73% for the samples for whichpercent activation could be determined. These data for the amount ofTGF-β in human serum can be compared with several previous reports. Avalue of 4.2±0.7 pmol/L (n=10) active TGF-β was obtained using the IL-4dependent HT-2 cell proliferation assay (Chao et al., Cytokine, 3, 292(1991)). However, when the serum was treated with acid, an increase ofgreater than 100-fold in TGF-β values was detected by the sameproliferation assay. This implies a mean value for activatable (i.e.,(α+1)) TGF-β of >420 pmol/L. In an earlier study (O'Connor-McCourt etal., supra.) using both a two-step competitive radio-receptor assay andthe NRK cell-soft agar growth system, it was reported that acid-ethanolextraction of serum (FCS, calf and human) gave (α+1) TGF-βconcentrations of 200-1000 pmol/L. A value for human serum for TGF-β1 of1,300 pmol/L and <5 pM for TGF-β2 measured by specific ELISAs has alsobeen reported (Dasch et al., Annals N.Y. Acad. Sci., 593, 303 (1990)).Of these data, only the low active TGF-β value of 4.2±0.7 pmol/L (n=10)differs substantially from the range of our ELISA values for human sera(Chao et al., supra).

Platelet-poor plasma samples were prepared from the same blood samplesused to prepare sera from the 4 donors in Table 4. There was nodifference within the accuracy of the assays between the amount of (α+1)TGF-β before or after acid treatment of the plasma samples, and afteracid treatment, the amount of active TGF-β was not significantlydifferent from the amount of (α+1) TGF-β (Table 6).

TABLE 6 Active and (α + 1) TGF-β concentrations in human platelet-poorplasma TGF-β (pmol/L) Unactivated plasma Acid-activated plasma DonorActive (α + 1) Active (α + 1) A <40 240 240 240 B 120 120 120 120 C 160320 320 320 D 200 240 240 280 Platelet-poor plasma were derived from thesame blood samples as the sera for Table 4 and were assayed in the sameexperiment for active and (α + 1) TGF-β by ELISA before and after acidactivation. All samples were determined in quadruplicate.

These data demonstrate that the plasma did not contain components whichinterfered with either ELISA, consistent with the finding for the seraderived from the same blood samples.

Comparison of the data in Tables 4 and 6 also shows that (α+1) TGF-βconcentrations and the proportions of TGF-β which were active were verysimilar in serum and platelet-poor plasma prepared from the same bloodsamples. These data implied that either the platelets had degranulatedto release their TGF-β during the preparation of the platelet-poorplasma so that the amounts of TGF-β were the same in plasma and inserum, or that platelet degranulation during clotting in the preparationof serum did not release active or latent TGF-β into the serum. Theserum and plasma TGF-β concentrations would then be similar because theserum and plasma did not contain a significant amount of active orlatent TGF-β from platelets which had degranulated after drawing theblood samples.

To examine whether the active or latent TGF-β in the serum and plasmasamples was derived from degranulation of platelets after drawing blood,(α+1) TGF-β concentrations in the sera, acid-extracted clots,platelet-poor plasma and platelets from seven donors were compared(Table 7).

TABLE 7 (α + 1) TGF-β concentrations in human serum, plasma, platelets,and acid-treated clots (α + 1) TGF-β (pmol/L) Platelet-poor Acid-treatedDonor Serum plasma Platelets clot E <40 40 1000 960 N 80 80 880 760 B120 120 1000 1200 D 280 280 1600 1600 A 320 360 1200 1200 C 440 440 1000720 M 1200 1400 760 760 Serum, platelet-poor plasma and platelets wereprepared from blood from 7 male donors. Clots were removed from theserum samples by centrifugation, washed, dissolved by acidification andneutralized. TGF-β was released from platelets by sonication whichlysed >90% of the platelets present. (α + 1) TGF-β in each sample wasassayed by ELISA in quadruplicate. TGF-β concentrations for plateletsand clots are calculated for the volume of blood from which they werederived.

The (α+1) TGF-β concentrations in serum and plasma derived from the sameblood samples were very similar, consistent with the data in Tables 4and 6. The average concentration of (α+1) TGF-β from the degranulatedplatelet samples was 1063 pmol/L and the average platelet concentrationby hemocytometer in the platelet preparations was 3.0×10¹¹/L, equivalentto an average of 2,100 molecules of TGF-β per platelet. This may becompared with a previous estimate of 500 to 2,000 molecules of TGF-β perplatelet recovered from “platelet secretate” (Wakefield et al., J. Biol.Chem., 263, 7646 (1988)). However, the surprising observation was thatthe (α+1) TGF-β concentrations of the degranulated platelets and theacid-extracted clots derived from the same blood samples were verysimilar. This observation implies that any active or latent TGF-βreleased by platelets which degranulated in the clots was almostentirely retained within the clot, since quantitative recovery of the(α+1) TGF-β was obtained from the clot after acid treatment. Theretention of (α+1) TGF-β in the clot would account for the closesimilarity of the (α+1) TGF-β concentrations in the sera and plasma andthis conclusion was tested further as described below. However, itshould be noted that the data do not preclude the possibility thatplatelets contain substantial amounts of latent TGF-β informs which arenot detected by the (α+1) TGF-β ELISA because they are not activated bythe defined acid-activation procedure.

No active TGF-β could be detected in the platelet releasate from freshlyprepared platelets, unlike the TGF-β obtained from blood bank platelets.When active recombinant human TGF-β1 was added to the platelet releasatecontaining the highest concentration of (α+1) TGF-β (1600 pmol/L) fromdonor D), the calibration curve for active TGF-β was superimposed on thecurve for the recombinant human TGF-β1 in TBS. These observations showthat the selectivity of the active TGF-β assay is at least 50-foldgreater for active TGF-β1 than latent TGF-β1.

The mean value for (α+1) TGF-β in platelet-poor plasma was 389±177pmol/L (n=7). Some of the reported values of TGF-β in platelet-poorplasma are similar to those described here. In two separate studiesusing acid-ethanol extraction of platelet-poor plasma and the MvLu cellbioassay, TGF-β concentrations of 212±132 pmol/L (n=9) and 244±40 pmol/L(range >80 to <400 pmol/L; n=10) were recently reported. Previously,Wakefield et al. (supra.) reported that human plasma containssignificant levels of TGF-β (60±24 pmol/L; n=10) and concluded thatlatent TGF-β does circulate in normal individuals (J. Clin. Invest., 86,1976 (1990)). One much lower value of 2.3 pmol/L (range 2.1 to 2.7pmol/L; n=9) for TGF-β1 in platelet-poor plasma assayed by a TGF-β1ELISA on acid-ethanol extracts has also been reported (Anderson et al.,Kidney International, 40, 1110 (1991)).

The similarity of both the (α+1) and active TGF-β concentrations inplatelet-poor plasma and serum from the same donor (Tables 4, 6, and 7)prompted the question of whether the TGF-β had been released by apartial degranulation of platelets when the blood samples were drawn andbefore the onset of clot formation in the serum samples. Since PDGF iscontained in the same platelet α-granules as latent TGF-β, a bioassayfor PDGF activity as a mitogen for human VSMCs was used to determine theextent of platelet degranulation during the preparation of theplatelet-poor plasma (Table 8).

TABLE 8 Mitogenic indices of human serum and plasma on human vascularsmooth muscle cells Mitogenic index Donor Serum Plasma B 45 0.7 H 52 1.4C 60 0.9 D 65 1.0 A 83 1.2

DMEM containing 5% serum or 20% platelet-poor plasma from five maledonors was added to quiescent, explant-derived human smooth muscle cellsand DNA synthesis was assayed in triplicate by incorporation of[³H]-thymidine between 12 hours and 36 hours after addition of thesamples. The mitogenic indices are the ratios of ³H counts incorporatedin the test cell samples to ³H counts in control cells treated withmedium alone (1,506±123 cpm). The mitogenic indices for the plasmasamples were unaffected by neutralizing antiserum to PDGF but werereduced by more than 52% for each of the serum samples.

Platelet-poor plasma had no significant mitogenic activity on humanVSMCs measured as a ratio of [³H]-thymidine incorporation in thepresence or absence of plasma (Table 8) and the ratio was unaffected byneutralizing antibody to PDGF. However, addition of 3.3 pmol/L PDGF tothe plasma samples caused an increase in the average mitogenic indexfrom 1.0 to 1.6 and this increase was blocked by neutralizing PDGFantibody. The platelet-poor plasma samples therefore contained less than3.3 pmol/L of active PDGF. In contrast, the human serum samples gavelarge mitogenic indices of 45 to 83 for the same cell preparation and atleast 52% of the mitogenic activity was reversed by neutralizingantibody to PDGF (50 mg/L).

This mitogenic activity attributable to PDGF is consistent with previousestimates that PDGF accounts for approximately 50% of platelet-derivedmitogenic activity of human serum, as assayed on glial cells orfibroblasts (Singh et al., J. Cell Biol., 95, 667 (1982)). The mitogenicstimulation reversible by neutralizing PDGF antibody (50 mg/L) in theserum samples corresponds to concentrations of human PDGF of greaterthan 300 pmol/L and less than 600 pmol/L in the human sera. This valuemay be compared with a reported concentration of PDGF in human serum of500 pmol/L by radio-receptor assay (Heldin et al., Exp. Cell. Res., 136,(1981)). A serum concentration of greater than 300 pmol/L thereforeimplies degranulation of most of the platelets during clot formation torelease PDGF into the serum under conditions in which the TGF-β remainsassociated with the clot. The undetectable PDGF activity in the plasmasamples indicates that the amount of PDGF in the plasma corresponds todegranulation of less than 5% of the platelets after bleeding.

Most previous work has shown that normal human plasma containsundetectable levels of PDGF. However, in one report (Heldin et al.,supra.), PDGF in human platelet-poor plasma was estimated at 33 pmol/Lby radio-receptor assay with a corresponding serum concentration of 500pmol/L. Thus, the preparation of platelet-poor plasma contained littleor no detectable PDGF from platelet degranulation during preparation inour experiments is consistent with previous data.

Taken together, these observations strongly imply (i) that the TGF-β inplatelet-poor plasma and serum do not result from platelet degranulationwhich occurs on or after taking the blood samples and (ii) that theconcentrations of (α+1) TGF-β in serum and plasma are very similarbecause platelet degranulation on clotting does not release (α+1) TGF-βinto the serum which can be detected by the (α+1) TGF-β assay. Similar(α+1) TGF-β concentrations in serum were obtained from repeated bleedsfrom the same donors. For example, donor A gave (α+1) TGF-βconcentrations of 240, 240, 320, 240, and 280 pmol/L from five bleeds atintervals of at least seven days. Furthermore, similar proportions of(α+1) TGF-β were active in repeated bleeds from the same donors. Theseobservations are consistent with negligible platelet degranulation afterthe blood samples are drawn since degranulation would be unlikely to besufficiently controlled to yield reproducible amounts of (α+1) TGF-β insera prepared from separate bleeds.

The data leave open the question of the origin of the TGF-β inplatelet-poor plasma. It is generally assumed that the plasma TGF-β ismainly derived from platelets and although plausible, this has not beendemonstrated experimentally. However, the ELISAs described here shouldfacilitate analysis of the mechanisms controlling platelet-poor plasmaconcentrations of active and (α+1) TGF-β. They should also allowexamination of correlations between TGF-β concentrations in plasma orserum and various diseases in which TGF-β may be implicated.

All publications, patents and patent applications are incorporatedherein by reference, except to the extent that the definitions in priorapplications and patents are inconsistent with the definitions herein.While in the foregoing specification this invention has been describedin relation to certain preferred embodiments thereof, and many detailshave been set forth for purposes of illustration, it will be apparent tothose skilled in the art that the invention is susceptible to additionalembodiments and that certain of the details described herein may bevaried considerably without departing from the basic principles of theinvention.

What is claimed is:
 1. An in vitro method for determining in bloodTGF-beta-1, TGF-beta-3, or a component that is bound by a TGF-beta-1ligand or a TGF-beta-3 ligand, said method comprising: (a) contacting amammalian blood-derived sample with a capture moiety capable of bindingto TGF-beta-1 or TGF-beta-3 to form a capture complex; (b) contactingthe capture complex with a detection moiety to form a detectablecomplex, wherein the detection moiety is capable of binding toTGF-beta-1 or TGF-beta-3 and comprises a detectable label or a sitewhich binds a detectable label, wherein the capture moiety or thedetection moiety comprises a TGF-beta type II receptor extracellulardomain; and (c) detecting the presence of the detectable complex,thereby identifying a mammal at risk for atherosclerosis or the effectof administering to a mammal a therapeutic agent which increases thelevel of TGF-beta-1, TGF-beta-3 or the component in said mammal.
 2. Atest kit for determining, in vitro, in a physiological sample obtainedfrom a mammal, TGF-beta-1, TGF-beta-3, or a component that is bound by aTGF-beta-1 ligand or a TGF-beta-3 ligand, comprising packaging materialenclosing, separately packaged, (a) a capture moiety capable of bindingTGF-beta-1, or TGF-beta-3; (b) a detection moiety capable of bindingTGF-beta-1 or TGF-beta-3, which moiety comprises a detectable label or abinding site for a detectable label, wherein either the capture moietyor the detection moiety comprises a TGF-beta type II receptorextracellular domain; and (c) instruction means directing the user tocorrelate the TGF-beta-1, TGF-beta-3, or component, in the sample withthe risk to said mammmal of atherosclerosis or with the effect of theadministration of a therapeutic agent which increases TGF-beta-1,TGF-beta-3 or the component in said mammal.
 3. An in vitro method fordetermining in blood TGF-beta-1, TGF-beta-3, or a component that isbound by a TGF-beta-1 ligand or a TGF-beta-3 ligand, consistingessentially of: (a) contacting a blood-derived sample from an individualwith a capture moiety capable of binding TGF-beta-1 or TGF-beta-3, toform a capture complex; (b) combining the capture complex with adetection moiety to form a detectable complex, wherein the detectionmoiety is capable of binding TGF-beta-1 or TGF-beta-3, and comprises adetectable label or a site which binds a detectable label, whereineither the capture moiety or the detection moiety comprises a TGF-betatype II receptor extracellular domain; and (c) determining the presenceof the detectable complex, so as to determine the presence ofTGF-beta-1, TGF-beta-3, or the component, in the sample.
 4. A test kitfor determining, in vitro, in a physiological sample, the level ofTGF-beta-1, TGF-beta-3, or a component that is bound by a TGF-beta-1ligand or a TGF-beta-3 ligand, comprising packaging material enclosing,separately packaged, (a) a capture moiety capable of binding TGF-beta-1or TGF-beta-3, and (b) a detection moiety capable of binding TGF-beta-1or TGF-beta-3, which moiety comprises a detectable label or a bindingsite for a detectable label, wherein either or both the capture moietyand the detection moiety is a fusion protein comprising the TGF-betatype II receptor extracellular domain.
 5. A method for determiningactive TGF-beta levels, comprising: (a) contacting a patient sample witha capture moiety that binds active TGF-beta-1 or active TGF-beta-1 andactive TGF-beta-3, to form a capture complex comprising said capturemoiety and active TGF-beta-1 or active TGF-beta-1 and active TGF-beta-3,wherein the capture moiety comprises a TGF-beta extracellular domaincomprising a signal peptide; (b) contacting the capture complex with adetection moiety which comprises a detectable label, or a site whichbinds a detectable label, to form a detectable complex; and (c)detecting the presence or amount of the detectable complex, so as todetermine the presence or amount of active TGF-beta-1 or activeTGF-beta-1 and active TGF-beta-3 in said sample.
 6. A method fordetermining in a patient sample the levels of active TGF-beta or acomponent that is bound by a TGF-beta-1 ligand or a TGF-beta-3 ligand,comprising: (a) contacting a patient sample with a capture moietycapable of binding active TGF-beta-1 or active TGF-beta-1 and activeTGF-beta-3, to form a capture complex; (b) contacting the capturecomplex with a detection moiety to form a detectable complex, whereinthe detection moiety comprises a detectable label or a site which bindsa detectable label; and (c) detecting the presence or amount of thedetectable complex, so as to determine the presence or amount of activeTGF-beta-1, active TGF-beta-1 and active TGF-beta-3, or the component,in said sample, wherein the presence or amount of active TGF-beta-1,active TGF-beta-1 and active TGF-beta-3, or the component, in saidsample is correlated to the presence or amount of active TGF-beta-1,active TGF-beta-1 and active TGF-beta-3, or the component, present invivo.
 7. A method for identifying a patient having, or at risk of, acondition associated with a TGF-beta deficiency or a deficiency in acomponent that binds a TGF-beta-1 ligand or a TGF-beta-3 ligand,comprising: (a) contacting a sample from said patient with a capturemoiety capable of binding TGF-beta-1 or TGF-beta-3 to form a capturecomplex; (b) contacting the capture complex with a detection moiety toform a detectable complex, which detection moiety comprises a detectablelabel or a site which binds a detectable label; and (c) detecting thepresence or amount of the detectable complex, so as to determine thepresence or amount of said TGF-beta-1, TGF-beta-3, or the component, insaid sample, thereby identifying a patient having or at risk of acondition associated with a TGF-beta deficiency or a deficiency in acomponent that binds a TGF-beta-1 ligand or a TGF-beta-3 ligand.
 8. Amethod for monitoring a mammal that has received one or moreadministrations of a therapeutic agent to increase the level of TGF-betaor a component that binds a TGF-beta-1 ligand or a TGF-beta-3 ligand,comprising: (a) contacting a biological sample from said mammal with acapture moiety capable of binding TGF-beta-1 or TGF-beta-3 to form acapture complex; (b) contacting the capture complex with a detectionmoiety to form a detectable complex, wherein the detection moiety whichcomprises a detectable label or a site which binds a detectable label;and (c) detecting the presence or amount of the detectable complex, soas to determine the presence or amount of said TGF-beta-1, TGF-beta-3,or the component, in said sample, thereby identifying the effect ofadministering to a mammal a therapeutic agent which increases the levelof said TGF-beta-1, TGF-beta-3, or the component in said mammal.
 9. Amethod for determining in a mammalian tissue sample in vitro TGF-beta-1,TGF-beta-3, or a component that is bound by a TGF-beta-1 ligand or aTGF-beta-3 ligand, said method comprising: (a) contacting a mammaliantissue sample with a TGF-beta type II extracellular domain to form acapture complex; and (b) detecting the presence of the capture complexTGF-beta-3, or the component in said tissue sample.
 10. A method fordetermining in a mammalian tissue sample TGF-beta-1, TGF-beta-3, or acomponent that is bound by a TGF-beta-1 ligand or a TGF-beta-3 ligand,comprising: (a) contacting a mammalian tissue sample with a capturemoiety capable of binding active TGF-beta-1 or active TGF-beta-3 to forma capture complex, wherein the capture moiety comprises a detectablelabel or a site which binds to a detectable label, and wherein thecapture moiety comprises a TGF-beta type II extracellular domain; and(b) detecting the presence or amount of the complex, so as to determinethe presence or amount of TGF-beta-1, TGF-beta-3, or the component insaid tissue sample.
 11. An in vitro method for determining in mammalianblood TGF-beta or a component that is bound by a TGF-beta-1 ligand or aTGF-beta-3 ligand, said method comprising: (a) contacting a mammalianblood-derived sample with a capture moiety, to form a capture complex;(b) contacting the capture complex with a detection moiety capable ofbinding TGF-beta to form a detectable complex, wherein either thecapture moiety or the detection moiety comprises the extracellulardomain of the TGF-beta type II receptor, and wherein the detectionmoiety comprises a detectable label or a site which binds a detectablelabel; and (c) detecting the presence of the detectable complex, so asto determine the presence of TGF-beta, or the component in said sample,thereby identifying a mammal at risk for atherosclerosis or the effectof administering to a mammal a therapeutic agent which increases thelevel of TGF-beta or the component in said mammal.
 12. A test kit fordetermining, in vitro, in a physiological sample obtained from a mammal,TGF-beta or a component that is bound by a TGF-beta ligand, comprisingpackaging material enclosing, separately packaged, (a) a capture moietycapable of binding TGF-beta; (b) a detection moiety capable of bindingTGF-beta, which moiety comprises a detectable label or a binding sitefor a detectable label, wherein either the capture moiety or thedetection moiety comprises the extracellular domain of the TGF-beta typeII receptor; and (c) instruction means directing the user to correlatethe TGF-beta or the component in the sample with the risk to said mammalof atherosclerosis or with the effect of the administration of atherapeutic agent which increases TGF-beta or the component in saidmammal.
 13. An in vitro method for determining in blood TGF-beta, or acomponent that is bound by a TGF-beta ligand, consisting essentially of:(a) contacting a blood-derived sample from an individual with a capturemoiety capable of binding TGF-beta to form a capture complex; (b)combining the capture complex with a detection moiety to form adetectable complex, wherein the detection moiety is capable of bindingTGF-beta, and comprises a detectable label or a site which binds adetectable label, wherein either the capture moiety or the detectionmoiety comprises the extracellular domain of the TGF-beta type IIreceptor; and (c) determining the presence of a detectable label in thedetectable complex, so as to determine the presence of TGF-beta or thecomponent in the sample.
 14. A test kit for determining TGF-beta levelsin vitro, comprising packaging material enclosing, separately packaged,(a) a capture moiety capable of binding TGF-beta and (b) a detectionmoiety capable of binding TGF-beta, which moiety comprises a detectablelabel or a binding site for a detectable label, wherein either thecapture moiety or the detection moiety is a fusion protein comprisingthe TGF-beta type II receptor extracellular domain.
 15. A method fordetermining active TGF-beta levels, comprising: (a) contacting a patientsample with a capture moiety that binds active TGF-beta to form acapture complex comprising said capture moiety and active TGF-beta,wherein the capture moiety comprises a TGF-beta type II extracellulardomain comprising a signal peptide; (b) contacting the capture complexwith a detection moiety to form a detectable complex, wherein thedetection moiety binds the capture complex and comprises a detectablelabel or a site which binds a detectable label; and (c) detecting thepresence or amount of the detectable complex, so as to determine thepresence or amount of active TGF-beta in said sample.
 16. A method fordetermining in a patient sample the levels of active TGF-beta or acomponent that is bound by a TGF-beta ligand, comprising: (a) contactinga patient sample with a capture moiety capable of binding activeTGF-beta to form a capture complex; (b) contacting the capture complexwith a detection moiety to form a detectable complex, wherein thedetection moiety binds the capture complex and comprises a detectablelabel or a site which binds a detectable label, wherein either thecapture moiety or the detection moiety comprises the extracellulardomain of the TGF-beta type II receptor; (c) detecting the presence oramount of the detectable complex, so as to determine the presence oramount of active TGF-beta or the component, in said sample is correlatedto the presence or amount of active TGF-beta or the component, presentin vivo.
 17. A method for identifying a patient having, or at risk of, acondition associated with a TGF-beta deficiency or a deficiency in acomponent that is bound by a TGF-beta ligand, comprising: (a) contactinga sample from said patient with a capture moiety capable of bindingTGF-beta to form a capture complex; (b) contacting the capture complexwith a detection moiety to form a detectable complex, wherein thedetection moiety binds the capture complex and comprises a detectablelabel or a site which binds a detectable label, wherein either thecapture moiety or the detection moiety comprises the extracellulardomain of the TGF-beta type II receptor; and (c) detecting the presenceor amount of the detectable complex, so as to determine the presence oramount of said TGF-beta or the component in said sample, therebyidentifying a patient having or at risk of a condition associated with aTGF-beta deficiency or a deficiency in the component.
 18. A method formonitoring a mammal that has received one or more administrations of atherapeutic agent to increase the level of TGF-beta or a component thatis bound by a TGF-beta ligand, comprising: (a) contacting a biologicalsample from said mammal with a capture moiety capable of bindingTGF-beta to form a capture complex; (b) contacting the capture complexwith a detection moiety to form a detectable complex, wherein either thecapture moiety or the detection moiety comprises the extracellulardomain of the TGF-beta type II receptor, and wherein the detectionmoiety binds the capture complex and comprises a detectable label or asite which binds a detectable label; and (c) detecting the presence oramount of the detectable complex, so as to determine the presence oramount of said TGF-beta or the component in said sample, therebyidentifying the effect of administering to said mammal a therapeuticagent which increases the level of said TGF-beta or the component insaid mammal.
 19. The method of claim 1 wherein the capture moiety isimmobilized on a solid substrate.
 20. The method of claim 1 wherein thecapture moiety is a solution phase capture moiety.
 21. The method ofclaim 1, 7 or 8 wherein the detection moiety is capable of bindinglatent and active TGF-beta-1 or latent and active TGF-beta-3.
 22. Themethod of claim 1 wherein the moiety that is not the TGF-beta type IIreceptor extracellular domain is an anti-TGF-beta antibody.
 23. Themethod of claim 1, 7 or 8 wherein the capture moiety is TGF-beta type IIreceptor extracellular domain and the detection moiety is ananti-TGF-beta antibody.
 24. The method of claim 1 wherein the presenceof the detectable complex is detected by reacting the detectable complexwith an antibody comprising a detectable label, which binds to saiddetectable complex, and determining the presence of the label.
 25. Themethod of claim 1 wherein the capture or the detection moiety comprisesa fusion protein comprising the TGF-beta type II extracellular domain.26. The method of claim 25 wherein the TGF-beta type II extracellulardomain has a methionine residue at position
 5. 27. The method of claim 1wherein either the capture moiety or the detection moiety is a fusionprotein comprising the TGF-beta Type II extracellular domain.
 28. Themethod of claim 27 wherein fusion protein is a prokaryotic fusionprotein.
 29. The method of claim 1 wherein either or both the capturemoiety and the detection moiety bind active TGF-beta-1, activeTGF-beta-3, or the component, but not latent TGF-beta-1 or latentTGF-beta-3.
 30. The method of claim 29 wherein the presence of activeTGF-beta-1, active TGF-beta-3, or the component, identifies a mammal atrisk for atherosclerosis or monitors the effect of administering to amammal a therapeutic agent which increases the level of TGF-beta-1,TGF-beta-3 or the component in said mammal.
 31. The test kit of claim 2wherein said capture moiety is immobilized on a solid substrate.
 32. Thetest kit of claim 2 wherein said capture moiety is present in solution.33. The test kit of claim 2 wherein one of the moieties is an antibody.34. The test kit of claim 2 wherein the capture moiety is a TGF-betatype II receptor extracellular domain.
 35. The test kit of claim 2wherein the TGF-beta type II receptor extracellular domain is derivedfrom a bacterial expression system.
 36. The test kit of claim 34 whereinthe detection moiety is an antibody.
 37. The test kit of claim 36further comprising, separately packaged, an antibody which binds to saiddetection moiety, which comprises a detectable label.
 38. The kit ofclaim 34 wherein the capture moiety is a fusion protein comprising theTGF-beta Type II extracellular domain.
 39. The kit of claim 38 whereinthe fusion protein is a prokaryotic fusion protein.
 40. The kit of claim2 wherein either or both the capture moiety and the detection moietybind active TGF-beta-1, active TGF-beta-3, or the component, but not thelatent TGF-beta-1 or latent TGF-beta-3.
 41. The kit of claim 40 whereinthe instruction means directs the user to correlate the level of activeTGF-beta-1, active TGF-beta-3, or the component, in the sample with therisk to said mammal of atherosclerosis or with the effect of theadministration of a therapeutic agent which increases the level ofTGF-beta-1, TGF-beta-3 or the component, in said mammal.
 42. The methodof claim 3 wherein the detection moiety is an antibody.
 43. The methodof claim 3 wherein the capture moiety is an antibody.
 44. The method ofclaim 3 wherein either the capture moiety or the detection moiety is afusion protein comprising the TGF-beta type II receptor extracellulardomain.
 45. The method of claim 44 wherein the TGF-beta type II receptorextracellular domain has a methionine residue at position
 5. 46. Themethod of claim 1 or 3 wherein the blood-derived sample is serum orplasma.
 47. The method of claim 3 wherein either or both the capturemoiety and the detection moiety bind active TGF-beta-1, activeTGF-beta-3, or the component, but not latent TGF-beta-1 or latentTGF-beta-3.
 48. The kit of claim 4 wherein the TGF-beta type II receptorextracellular domain has a methionine residue at position
 5. 49. The kitof claim 4 wherein fusion protein is a prokaryotic fusion protein. 50.The method of claim 5, 6, 7 or 8 wherein the moiety which comprises theTGF-beta type II extracellular domain is a fusion protein comprising theTGF-beta extracellular domain.
 51. The method of claim 5, 6, 7 or 8wherein the sample is a blood-derived sample.
 52. The method of claim 51wherein the sample is serum or plasma.
 53. The method of claim 51wherein the sample is not acid activated prior to step (a).
 54. Themethod of claim 6, 7, or 8 wherein in either or both the capture anddetection moieties bind active but not latent TGF-beta-1, the component,or active but not latent TGF-beta-3.
 55. The method of claim 54 whereinthe presence or amount of active TGF-beta-1, the component, or activeTGF-beta-1 and active TGF-beta-3 is detected.
 56. The method of claim 55further comprising correlating the presence or amount of activeTGF-beta-1, active TGF-beta-1 and active TGF-beta-3, or the component,in said sample to the presence or amount of the active form ofTGF-beta-1, the active form of TGF-beta-1 and the active form ofTGF-beta-3, or the component, present in vivo.
 57. The method of claim 7or 8 wherein the capture or detection moiety comprises a TGF-betaextracellular domain comprising a signal peptide.
 58. The method ofclaim 7 wherein the condition is a vascular condition.
 59. The method ofclaim 7 wherein the condition atherosclerosis.
 60. The method of claim 7or 8 wherein the capture or detection moiety comprises a TGF-beta typeII extracellular domain and wherein the moiety that is not the TGF-betatype II receptor extracellular domain is an anti-TGF-beta antibody. 61.The method of claim 9 wherein the tissue sample is part of a mammalianblood vessel.
 62. The method of claim 61 wherein the vessel is theaorta.
 63. The method of claim 9 wherein step (b) identifies a mammal atrisk of a condition associated with TGF-beta deficiency.
 64. The methodof claim 9 wherein step (b) identifies the effect of administering to amammal a therapeutic agent which increases the level of TGF-beta-1 orTGF-beta-3 in said mammal.
 65. The method of claim 9 wherein the capturecomplex is detected by a reaction with a labeled antibody.
 66. Themethod of claim 9 or 10 wherein the capture moiety is TGF-beta type IIextracellular domain.
 67. The method of claim 9 or 10 wherein thecapture moiety is a fusion protein comprising the TGF-beta type IIextracellular domain.
 68. The method of claim 66 wherein the TGF-betatype II receptor extracellular domain has a methionine residue atposition
 5. 69. The method of claim 10 wherein the capture moietycomprises a detectable label.
 70. The method of claim 69 wherein thedetectable label is a fluorescent label.
 71. The method of claim 10wherein the presence of TGF-beta-1, TGF-beta-3, or the component,identifies a mammal at risk for a condition associated with a TGF-betadeficiency or monitors the effect of administering to a mammal atherapeutic agent which increases the level of TGF-beta or the componentin said mammal.
 72. The method of claim 71 wherein the condition is avascular condition.
 73. The method of claim 71 wherein the condition isatherosclerosis.
 74. The method of claim 8 wherein the presence oramount of the detectable complex after administration is compared to thepresence of the complex or the amount of the complex formation prior tosaid administration.